LPS Priming of Stromal Cells to Generate LPS-Specific Exosome Educated Macrophages

ABSTRACT

The disclosure relates to an ex vivo generated population of educated macrophages specific to LPS and methods of making and using such macrophages.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application claims priority to U.S. Provisional Application No.62/629,479, filed Feb. 12, 2018, which is incorporated herein byreference in its entirety.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH

This invention was made with government support under CA014520 awardedby the National Institutes of Health. The government has certain rightsin the invention.

BACKGROUND

Radiation, delivered therapeutically, accidentally, or maliciously, canlead to an acute radiation syndrome (ARS) with life threateningtoxicities. High-dose radiation causes damage to highly proliferativecells such as those found in the bone marrow, GI-tract and skin. Currentstandard of care involves supporting victims with antibiotics andtransfusions until they can undergo an allogeneic bone marrow transplant(BMT) from a suitable donor. Unfortunately, the entire BMT process canoften take weeks to identify and collect cells from a donor, and aredifficult to perform on a large-scale in the event of a widespreadexposure. Moreover, allogeneic BMTs have their own set of complications,including engraftment failure, opportunistic infections and/orgraft-versus-host-disease (GVHD), making them potentially as toxic asthe radiation injury itself. Consequently, adjunct therapies havereceived special attention for treatment of radiation injury, but todate, the only approved treatment agents are colony-stimulating factorssuch as G-CSF. Unfortunately, for these factors to be effective thepatient's own hematopoietic stem cells have to be spared from radiationeffects.

Attractive alternatives to treat ARS focus on allogeneic,“off-the-shelf” cell-based therapies that can accelerate the repair oftissue injury after radiation without relying on the patient's ownremaining healthy cells. Ideally, cell-based therapies can becryopreserved and ready for a quick infusion without extensive tissuematching to the recipient. Among the stromal cells currently exploredare multipotent mesenchymal stem cells (MSCs) derived from the bonemarrow (BM) or tissue fibroblasts. MSCs are capable of self-renewal anddifferentiation into osteocytes, chondrocytes and adipocytes, makingthem attractive candidates to treat tissue injury. MSCs have strongimmunosuppressive properties and can control inflammation by modifyingthe proliferation and cytokine production of immune cells. Fibroblastsare also useful to treat tissue injury and are very similar to MSCs inmany properties such as morphology, surface marker profile and abilityto differentiate into other tissues. (Denu et al. Acta Haematol. 2016;136(2):85-97) MSCs have shown promise in preclinical studies in rodentmodels of radiation injury and have spurred human clinical trials fortreating autoimmune and degenerative diseases. However, whiletherapeutic MSCs show promise, they often fail to demonstrate clearefficacy in many clinical trials, and have not yet been approved totreat ARS.

While studies indicate that MSCs promote tissue repair based on theirdifferentiation potential, the lack of correlation between cellengraftment and differentiation at the site of injury with functionalimprovement suggest that MSCs achieve in vivo therapeutic effects bycommunicating with other effector cells. MSCs are thought to exerttherapeutic effects via antigen-presenting cells (APCs) such asmonocytes and macrophages. Macrophages can polarize generally into twobroad phenotypes: classically activated (M1) macrophages, which mediatetissue damage and are considered “pro-inflammatory”, or alternativelyactivated (M2) macrophages, which contribute to wound healing and tissuerepair and are “anti-inflammatory”. Direct co-culture of MSCs withmacrophages educates the macrophages to MSC-educated macrophages (MEMs)that increase the expression of specific surface markers (CD206) andcytokines (IL-6 and IL-10). The therapeutic utility of macrophageeducation by MSCs was demonstrated by enhanced survival from lethalradiation injury using a xenogeneic mouse model treated with MEMs ascompared to infusions of MSCs or macrophages alone.

While advancements have been made in the treatment of acute radiationsyndrome and other diseases using MSCs and MEMs, a need exists in theart for further development of new treatment methods and compositionsutilizing both MSCs and alternatively activated educated macrophages.

SUMMARY OF THE INVENTION

In a first aspect, provided herein is a method for generating aneducated macrophage, the method comprising the steps of isolating anextracellular vesicle from a mesenchymal stromal cell previously exposedto lipopolysaccharide (LPS), and co-culturing a CD14+ cell with theextracellular vesicle in vitro until the CD14+ cell acquires ananti-inflammatory macrophage phenotype. In some embodiments, the CD14+cell and the extracellular vesicle are co-cultured for at least 2 days.In some embodiments, the mesenchymal stromal cell is exposed to LPS forat least 2 hours. In some embodiments, the mesenchymal stromal cell isexposed to about 50 ng/ml to about 200 ng/ml LPS. In some embodiments,the mesenchymal stromal cell is exposed to about 800 ng/ml to about 1200ng/ml LPS. In some embodiments, the mesenchymal stromal cell is amesenchymal stem cell. In some embodiments, the mesenchymal stromal cellis a fibroblast. In some embodiments, the CD14+ cell is a macrophage. Insome embodiments, the CD14+ cell is a monocyte and wherein the CD14+monocyte and the extracellular vesicle are co-cultured for at least 5days.

In a second aspect, provided herein is a population of anti-inflammatorymacrophages produced by the methods described herein wherein themesenchymal stromal cell is exposed to about 50 ng/ml to about 200 ng/mlLPS, wherein the anti-inflammatory macrophage phenotype is characterizedas CD206 high, PD-L1 high, PD-L2 high, CD16 high and CD73 high comparedto control macrophages.

In a third aspect, provided herein is a population of anti-inflammatorymacrophages produced by the methods described herein wherein themesenchymal stromal cell is exposed to about 800 ng/ml to about 1200ng/ml LPS, wherein the anti-inflammatory macrophage phenotype ischaracterized as FLT-3L high, IL-15 high, CD73 high, CD86 low, andHLA-DR low as compared to control macrophages.

In a forth aspect, provided herein is a method for generating aneducated monocyte, the method comprising the steps of isolating anextracellular vesicle from a mesenchymal stromal cell previously exposedto lipopolysaccharide (LPS), and co-culturing a CD14+ monocyte with theextracellular vesicle in vitro until the CD14+ monocyte acquires ananti-inflammatory monocyte phenotype. In some embodiments, the CD14+monocyte and the extracellular vesicle are co-cultured for at least 2hours. In some embodiments, the CD14+ monocyte and the extracellularvesicle are co-cultured for at least 24 hours. In some embodiments, themesenchymal stromal cell is exposed to LPS for at least 12 hours. Insome embodiments, the mesenchymal stromal cell is exposed to about 50ng/ml to about 200 ng/ml LPS. In some embodiments, the mesenchymalstromal cell is exposed to about 800 ng/ml to about 1200 ng/ml LPS. Insome embodiments, the mesenchymal stromal cell is a mesenchymal stemcell. In some embodiments, the mesenchymal stromal cell is a fibroblast.

In a fifth aspect, provided herein is a population of anti-inflammatorymonocytes produced by the method of claim 17, wherein theanti-inflammatory monocyte phenotype is characterized as PD-L1 high,CD206 low, CD163 low, IL-15 high, CD73 high, CD86 low, CD16 low and IL-6high as compared to control monocytes.

INCORPORATION BY REFERENCE

All publications, patents, and patent applications mentioned in thisspecification are herein incorporated by reference to the same extent asif each individual publication, patent, and patent application wasspecifically and individually indicated to be incorporated by reference.

BRIEF DESCRIPTION OF DRAWINGS

The patent or patent application file contains at least one drawing incolor. Copies of this patent or patent application publication withcolor drawings will be provided by the Office upon request and paymentof the necessary fee.

The invention will be better understood and features, aspects, andadvantages other than those set forth above will become apparent whenconsideration is given to the following detailed description thereof.Such detailed description makes reference to the following drawings.

FIGS. 1A-1D demonstrate that extracellular vesicles (EVs) isolated frommesenchymal stem cells (MSCs), include small exosomes (50-250 nm) andlarger micro-vesicles (500-1000 nm), and exosome-sized EVs predominateamong the isolated EVs. EVs were isolated from bone marrow mesenchymalstem cells (BM-MSC) as described in the Examples, and EVs were analyzedby TEM and two different instruments to quantify the mean and mode ofthe particle diameter and particle concentration. (A and B) TEM of twodifferent preparations indicated that the particles had the typicalcup-shaped vesicular appearance of EVs and generally measured less than250 nm. (C) The preparations characterized by dynamic light scatteringusing the Nanosight NS300 indicated that the majority of particles were95 nm with a range of 50 to 250 nm. (D) The preparations characterizedby resistive pulse sensing using the qNano Nanoparticle instrumentmatched the characterization obtained from the Nanosight NS300 and alsoindicated that the majority of the particles are in the 50-250 nm range.Based on these characterizations the majority of the particles wereprimarily exosomes-sized EVs.

FIG. 2 shows the surface marker profile of the EVs from unstimulatedMSCs (MSC-EVs) and LPS-high stimulated MSCs (LPS-high-EVs). The resultsshown represent samples from two different human MSC donors. A total of37 known EV surface markers were examined. Surface markers positivelyidentified on the EVs are expressed as mean fluorescence intensity(MEI)±SEM.

FIGS. 3A-3B show surface marker profiles of (A) mean fluorescentintensity (MFI) and (B) % cells respectively, generated by flowcytometry for control macrophages, MSC exosome educated macrophages(EEMs), LPS-low exosome educated macrophages (LPS-low-EEMs), andLPS-high exosome educated macrophages (LPS-high-EEM). The results shownrepresent samples from at least three human donors wherein MSCs weremobilized using G-CSF. The percent (%) of CD14⁺ cells positive (+/−SEM)for each recited marker were measured by flow cytometry. Macrophageswere cultured for 7 days (Day 7 macrophages) followed by an additional 3days of unstimulated culture (control macrophages) or 3 days ofco-culture with either exosomes from MSCs (EEM), exosomes from MSCsprimed with low concentration LPS (LPS-low-EEMs), or exosomes from MSCsprimed with high concentration LPS (LPS-high-EEMs). Macrophages at day10 of culture (Day 10 macrophages) were used for flow cytometry. Pvalues were compared to control macrophages (along the x-axis) or withingroups (bars); *p</=0.05, **p</=0.01, ***p</=0.001, ****p</=0.0001.

FIG. 4 shows surface marker profiles as a percentage of cells generatedby flow cytometry for control monocytes, MSC exosome educated monocytes(EEMos), and LPS-high exosome educated monocytes (LPS-high-EEMos).Frozen stocks of monocytes from at least three different human donorsmobilized with G-CSF were used. The monocytes were thawed and placed inculture media and either left unstimulated (control monocytes) ordirectly educated with exosomes for approximately 24 hours with eitherexosomes from MSCs (EEMo) or exosomes from MSCs primed with highconcentration LPS to generate LPS-high-EEMos. P values were compared tocontrol macrophages (along the x-axis) or within groups (bars);*p</=0.05, **p</=0.01, ***p</=0.001, ****p<=/=0.0001.

FIGS. 5A-5C show gene expression by RT-PCR of monocytes from multiplehuman isolates which were either unstimulated (control) or educated for24 hours with exosomes from MSCs to produce EEMos and LP-high-EEMos asdescribed in Example 1. After education, cells were collected, RNAisolated and analyzed by RT-PCR for gene expression. The fold change ofgene expression is normalized to the expression level of the GAPDHhouse-keeping gene in unstimulated control macrophages and set at avalue of 1.0. FIG. 5A compares the gene expression of IL-6. FIG. 5Bcompares the gene expression of IL-8, IDO and FGF2. FIG. 5C compares thegene expression of IL-15, IL-10, IL-12, VEGF-A, EGF and IL-7. P valueswere determined compared to controls; *p</=0.05, **p</=0.01,***p</=0.001, ****p</=0.0001.

FIG. 6 shows that LPS-high-EEMs are strongly phagocytic using pHrodoGreen E. coli bioparticles. Macrophages at day 7 of culture (Day 7macrophages) were tested in their unstimulated state (control) orco-culture for an additional 3 days with either M1 promoting factors (M1stimulation), exosomes from MSCs (EEMs), or exosomes from MCSs primedwith a low (LPS-low-EEMs) or high (LPS-high-EEMs) concentration of LPSas described in methods. Post co-culture Day 10 macrophages were treatedwith pHrodo Green E. coli bioparticles and the ratio of CD14+ cellspositive for pHrodo Green E. coli bioparticles to total CD14+ cells wasdesignated as percent (%) cells as determined by flow cytometry. Pvalues were compared to control; *p</=0.05, **p</=0.005, ***p</=0.0005.

FIGS. 7A-7B show Immuno-potency assay (IPA) for growth inhibition ofT-cells and measures the % proliferation of antibody activated (A)CD4+(helper T cells) or (B) CD8+(cytotoxic T cells) when co-cultured atvarious ratios with test cells that were either MSCs, controlmacrophages, EEMs, or LPS-EEMs. The % cell proliferation is compared toactivated cells without the addition of test cells.

FIGS. 8A-8C show that treatment with LPS-high-EEMs significantlyincreased survival, improved weight loss and clinical scores in miceafter challenge with lethal radiation. (A-C) On day 0,NOD.Cg-Prkdc^(scid)Il2rg^(imIWjl)/SzJ (NSG) mice (from The JacksonLaboratory) received 4 gray (Gy) of lethal radiation followed by anintravenous treatment 4 hours later with PBS, 1×10⁶ human bone marrowMSCs, Day 10 control macrophages, EEMs, LPS-low-EEMs or LPS-high-EEMsgenerated as described in the Examples. FIG. 5A shows a graph of thesurvival curve vs days post-challenge compared by log rank analysis. Pvalue comparing LPS-high-EEMs to the other groups was >0.0001. (B) Mean% weight change vs days post-challenge compared with Day 0 for eachgroup. P value comparing LPS-high-EEMs to the other groups was >0.0001.(C) Overall clinical score (weight loss, posture, activity and furtexture) vs days post-challenge. P value comparing LPS-high EEMs to theother groups was >0.0001.

FIGS. 9A-9B show tissue histology of bone marrow and spleen samples.Histology shows that human LPS-high-EEM treatment protects againsttissue damage in the bone marrow and spleen of mice after lethalradiation injury. On day 0, MSG mice received either no radiation(normal healthy control) or 4 Gy of lethal radiation. 4 hours afterexposure, mice received an intravenous treatment of PBS or 10LPS-high-EEMs. Bone marrow and spleen tissue samples were taken 9 dayspost radiation from PBS controls and LPS-high-EEM treated mice. Sampleswere also taken from the LPS-high-EEM treated mice 31 days and 53 dayspost radiation exposure. FIG. 9A shows 20× images of H&E stained femoralbone marrow sections from each group. FIG. 9B shows 20× images of H&Estained spleen sections.

FIGS. 10A-10C show treatment with LPS-high-EEMos significantly increasedsurvival, improved weight loss and clinical scores in mice afterchallenge with lethal radiation. (A-C) On day 0, NOD.Cg-PrkdcscidIl2rgmiWJlfSzJ/(NSG) mice (from The Jackson Laboratory) received 4 gray(Gy) of lethal radiation. 4 hours later mice received an intravenoustreatment with either PBS or 1×10⁷ of control monocytes, EEMos, orLPS-high-EEMos generated as described in Example 1. FIG. 10A shows agraph of the survival curve vs days post-challenge compared by log rankanalysis. P value comparing LPS-high-EEMos to the other groupswas >0.0001. FIG. 10B shows overall clinical score (weight loss,posture, activity and fur texture) vs days post-challenge. FIG. 10Cshows mean percent weight change vs days post-challenge compared withDay 0 for each group. P value comparing LPS-high-EEMs to the othergroups was >0.0001.

FIG. 11 shows direct treatment with exosomes is ineffective at treatingmice after challenge with lethal radiation. The exosome treatment dosage(particle number) used to treat each mouse was the same dose used inco-culture with CD14+ macrophages or monocytes to create LPS-high-EEMsor LPS-low-EEMos. On day 0, NSG mice (from the Jackson Laboratory)received 4 Gy of lethal radiation. 4 hours after radiation exposure,mice received intravenous treatment with either PBS, exosomes isolatedfrom MSCs, or exosomes isolated from MSCs primed with LPS at aconcentration of 2.5×10⁹ particles/100 μl PBS.

FIGS. 12A-12C show that direct treatment with high doses of exosomesisolated from LPS primed MSCs can significantly increase survival,decrease radiation exposure associated weight loss, and improve clinicalscores in mice after challenge with lethal radiation. On day 0, NSG mice(from The Jackson Laboratory) received 4 Gy of lethal radiation. 4 hoursafter radiation exposure, mice received intravenous treatment witheither PBS, exosomes isolated from unstimulated MSCs (MSC-EVs at aconcentration of 5.0×10⁹ particles/100 μl PBS), exosomes isolated fromMSCs primed with high concentration LPS (LPS-high-EVs at a concentrationof 5.0×10⁹ particles/100 μl PBS), or monocytes educated with exosomes ata concentration 5.0×10⁹ particles/100 μl, the exosomes for monocyteeducation having been isolated from MSCs primed with high concentrationLPS. FIG. 12A shows a graph of the survival curve vs days post-challengecompared by log rank analysis. P value comparing treatment withLPS-high-EEMos, MSC-EVs, and LPS-high-EVs to PBS control was >0.001and >0.01 respectively. FIG. 12B shows overall clinical score (weightloss, posture, activity and fur texture) vs days post-challenge. P valuecomparing treatment with LPS-high-EEMos to PBS control was >0.01. FIG.12C shows mean percent weight change vs days post-challenge comparedwith Day 0 for each group.

DETAILED DESCRIPTION OF THE INVENTION

The present disclosure broadly relates to an educated CD14⁺ cell(macrophage or monocyte) as well as methods for making and using such acell. The educated CD14⁺ cell may be a cell with an anti-inflammatory,immunosuppressive, tissue reparative phenotype. Methods of the presentinvention broadly relate to derivation of extracellular vesicles fromLPS-treated mesenchymal stromal cells (LPS-EVs) and their use in aco-culture with CD14+ cells to generate LPS-specific educatedmacrophages (LPS-EEMs) or LPS-specific educated monocytes (LPS-EEMos).The disclosure also broadly relates to methods of treatment usingLPS-EVs, LPS-EEMS, or LPS-EEMos.

In one aspect of the invention, mesenchymal stromal cells are culturedin the presence of LPS to generate LPS-primed stromal cells.Extracellular vesicles isolated from the LPS-primed stromal cells(LPS-EVs) are co-cultured with CD14⁺ monocytes or macrophages to yieldeducated macrophages (LPS-EEMs) or educated monocytes (LPS-EEMos) with acharacteristic cytokine profile, expression profile and phenotype asdescribed herein. LPS-EVs, educated macrophages, or educated monocytesgenerated by the methods of the present invention may be used to treator prevent a disease by administering the educated cells to a subject inneed thereof.

As used herein, “educated macrophage” refers to a LPS-specificanti-inflammatory, tissue reparative, immunosuppressive macrophagegenerated ex vivo by co-culturing a CD14⁺ monocyte or macrophage with anextracellular vesicle obtained from a LPS-treated mesenchymal stromalcell. In one embodiment, the educated macrophages are anti-inflammatory,immunosuppressive, and tissue reparative macrophages generated byco-culturing CD14⁺ monocytes or macrophages with extracellular vesiclesderived from LPS-primed MSCs. In one embodiment, the educatedmacrophages are anti-inflammatory, immunosuppressive, and tissuereparative macrophages generated by co-culturing CD14⁺ monocytes ormacrophages with extracellular vesicles derived from LPS-primedfibroblasts.

As used herein, “educated monocyte” refers to a LPS-specificanti-inflammatory, tissue reparative, immunosuppressive monocytegenerated ex vivo by co-culturing a CD14⁺ monocyte with an extracellularvesicle obtained from a LPS-treated mesenchymal stromal cell. In oneembodiment, the educated monocytes are anti-inflammatory,immunosuppressive, and tissue reparative monocytes generated byco-culturing CD14⁺ monocytes with extracellular vesicles derived fromLPS-primed MSCs. In one embodiment, the educated monocytes areanti-inflammatory, immunosuppressive, and tissue reparative monocytesgenerated by co-culturing CD14⁺ monocytes with extracellular vesiclesderived from LPS-primed fibroblasts.

Co-Culture

CD14⁺ cells are co-cultured with LPS-EVs to yield LPS-specific educatedmacrophages (LPS-EEMs) or LPS-specific educated monocytes (LPS-EEMos).Methods of co-culturing CD14⁺ cells with mesenchymal stem cells (MSCs),mesenchymal stromal cells, or tissue-specific extracellular vesicles(EVs) to generate MSC-educated macrophages (referred to herein asBM-MEM) or exosome educated macrophages (EEMs), respectively, have beendescribed, see U.S. Pat. No. 8,647,678 and U.S. Patent Publication No.2016/0082042, each of which is incorporated herein by reference.

CD14⁺ cells are co-cultured ex vivo with LPS-EVs in any culture mediumknown in the art suitable for survival and growth of the co-culturecomponents. CD14⁺ cells may be co-cultured in culture plates, cultureflasks or in hollow fiber systems. To generate educated monocytes, theco-cultures may be maintained for between 2 hours and 5 days.Co-cultures may generate educated monocytes with the desiredimmune-phenotype after 2 hours, 3 hours, 4 hours, 5 hours, 8 hours, 10hours, 12 hours, 15 hours, 18 hours, 20 hours, 24 hours, 36 hours, 48hours, 50 hours, 72 hours, 4 days, or 5 days. In some embodiments,co-cultures yield educated monocytes after 24 hours. In someembodiments, co-cultures yield educated monocytes after 48 hours. Togenerate educated macrophages, the co-cultures may be maintained forbetween 1-20 days. Co-cultures may generate educated macrophages withthe desired immuno-phenotype after 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11,12, 13, 14, or more than 15 days. In some embodiments, co-cultures yieldeducated macrophages after 10 days. In some embodiments, co-culturesyield educated macrophages after 5 days. In some embodiments,co-cultures yield educated macrophages after 3 days. In one embodiment,co-cultures yield educated macrophages after 1 day. In general, toproduce a population of educated macrophages starting with a populationof CD14⁺ monocytes, cells are co-cultured with LPS-EVs for at least 5days to generate educated macrophages or CD14⁺ monocytes aredifferentiated to macrophages and the resulting CD14⁺ macrophages areco-cultured with LPS-EVs for at least one day to generate educatedmacrophages.

In some cases, LPS-EVs are subjected to additional purification stepsprior to use in co-culture. LPS-EVs can be added in a single dose orrepeated doses to CD14⁺ cultures to generate educated CD14⁺ cells. Inone embodiment, an additional centrifugation step is added to separateexosomes from micro-vesicles and other extra-cellular vesicles.

For co-cultures of the present invention, monocytes or macrophages canbe co-cultured with LPS-EVs such that the cells are in direct physicalcontact with the extracellular vesicles. Alternatively, the co-culturecomponents can be placed in sub-compartments that are in fluidcommunication but separated by a semi-permeable membrane. Thesemi-permeable membrane allows the exchange of soluble medium componentsand factors secreted by the cells but not the cells per se. The poreswithin the semi-permeable membrane are sufficiently small to preventcell penetration but large enough to allow soluble medium components topass across the membrane, and are typically are between 0.1-1.0 μm, butother pore sizes can be suitable.

Various methods of cell separation and isolation are known in the artand can be used to separate the educated macrophages or educatedmonocytes from the LPS-EVs depending on factors such as the desiredpurity of the isolated cell populations. Macrophages are stronglyadherent to solid culture surfaces and monocytes are weakly adherent toculture surfaces which may aid in separation and isolation of theeducated CD14⁺ cells. For example, educated macrophages or educatedmonocytes can be isolated from the co-culture using flow cytometry,magnetic-based sorting, scraping from plates, a digestive process (e.g.,trypsinization and EDTA), or low speed centrifugation. In someembodiments, educated macrophages or educated monocytes can be separatedfrom the LPS-EVs by removal of the culture medium containing the LPS-EVsfollowed by multiple washing steps. Educated macrophages can bemaintained in culture in any medium that supports macrophages in vitro.Educated monocytes can be maintained in culture in any medium thatsupports monocytes in vitro. Also, educated macrophages or educatedmonocytes can be stored using methods known in the art including, butnot limited to, refrigeration, cryopreservation, vitrification, andimmortalization.

As used herein, “CD14⁺ cell” refers to a monocyte or a macrophage. CD14⁺cells can be derived from any suitable source. The skilled artisan willappreciate the advantageous efficiency of generating macrophages fromperipheral blood monocytes for co-cultures. Alternatively, macrophagescan also be isolated from cellular outgrowth of a tissue sample takenfrom an individual or from pluripotent stem cells. Monocytes can becultured for various times and under various conditions beforeco-culture or can be added to the exosomes or extracellular matrixdirectly for co-cultures. In one embodiment, monocytes are harvestedfrom a subject by leukapheresis. In one embodiment, CD14+ cells areisolated from peripheral blood. In one embodiment, CD14+ cells areisolated from peripheral blood of a patient who has first been treatedwith an agent including but not limited to G-CSF, GM-CSF, Mozobil andthe like to mobilize cells into the peripheral blood. In one embodiment,CD14+ cells are isolated from peripheral blood with G-CSF stimulation.In one embodiment CD14+ cells are isolated from bone marrow aspirates.In one embodiment CD14+ cells are isolated from tissues or organs ofinterest. In one embodiment CD14+ cells are derived from pluripotentstem cells such as embryonic stem cells or induced pluripotent stemcells.

As used herein “macrophage” refers to a mononuclear phagocytecharacterized by the expression of CD14 and lack of expression ofdendritic or mesenchymal cell markers.

As used herein “mononuclear leukocytes” or “monocytes” are white bloodcells that can differentiate into macrophages when recruited to tissuesand can influence both innate and adaptive immune system.

As used herein, “high” means that the cells are characterized by higherexpression of a particular cytokine, chemokine, growth factor or cellsurface marker compared to control macrophages or monocytes culturedunder the same conditions without tissue-specific cells or extracellularfactors. Expression of markers may be measured by any means known in theart, including but not limited to, gene expression analysis (qPCR),Western blot, secretion product measurement by ELISA, multiplexdetection systems, transcriptome analysis or flow-cytometry. Forexample, “IL-6 high” indicates that macrophages co-cultured withtissue-specific cells or extracellular factors express higher amounts ofIL-6 than macrophages that have not been co-cultured withtissue-specific cells or extracellular factors. Similarly, “low” meansthat the cells are characterized by lower expression of a particularcytokine. For example, “IL-12 low” indicate that macrophages co-culturedwith tissue-specific cells or extracellular factors express loweramounts of IL-12 than macrophages that have not been co-cultured withtissue-specific cells or extracellular factors. “Low” can also mean thatthe expression levels or secretion levels are below the detection limit.

Primed Stromal Cells and Extracellular Vesicles

The skilled artisan will appreciate that monocytes, macrophages,mesenchymal stromal cells, fibroblasts, mesenchymal stem cells, andextracellular vesicles employed in methods described herein can becultured or co-cultured in any medium that supports their survival andgrowth. In some embodiments, the medium is a serum-free mediumsupplemented with chemically defined mammalian serum supplement. In someembodiments the medium is supplemented serum-free medium including butnot limited to X-VIVO™ 15, CTS™ STEMPRO™ MSC serum-free media (SFM), orSTEMPRO™-34 SFM. One may also use conventional culture media with serumor an animal supplement depleted of endogenous EVs which may be presentin the serum. EVs may be removed from the serum by means such asultracentrifugation or ultrafiltration. Suitable serum from whichendogenous EVs may be removed include but are not limited to fetalbovine serum, fetal calf serum, human serum, and human A/B serum. Forshort term cultivation of about 1 day to about 3 days, conventionalculture medium without serum has also been used. In one embodiment, themedium uses human platelet lysates to replace the human AB serum in theculture medium for macrophage and monocyte cultures. In someembodiments, in order to isolate EVs for the cells of interest, theculture medium is free of endogenous EVs present in either mammalianserum or protein supplements derived from humans or animals such ashuman platelet lysate. Mesenchymal stem cells, extracellular vesiclesand macrophages can be autologous, syngeneic, allogeneic, or third partywith respect to one another.

As used herein, “mesenchymal stromal cells” refers to mesenchymal stemcells (MSC) or fibroblasts.

As used herein, “mesenchymal stem cells (MSC)” refers to thefibroblast-like cells that reside within virtually all tissues of apostnatal individual. An ordinarily skilled artisan will appreciate thatthe cells referred to herein as mesenchymal stem cells are also known inthe art as mesenchymal stromal cells, marrow stromal cells, multipotentstromal cells, and other names. An MSC within the scope of thisdisclosure is any cell that can differentiate into osteoblasts,chondrocytes, myocytes, and adipocytes. An MSC within the scope of thisdisclosure is positive for the expression of CD105, CD73, and CD90 whilelacking expression of CD45, CD34, CD14 or CD11b, CD79a or CD19, andHLA-DR surface molecules. (Dominici et al. Minimal criteria for definingmultipotent mesenchymal stromal cells. The International Society forCellular Therapy position statement, (2006), Cytotherapy, 8(4):315-317).While these markers are known to characterize MSCs derived from mosttissues, it is understood in the art that MSCs from some sources couldexhibit differences in cell surface marker expression. Within bonemarrow, MSCs provide the stromal support tissue for hematopoietic stemcells. MSCs can differentiate into cells of the mesenchymal lineage. Insome embodiments, MSCs are co-cultured with CD14⁺ cells to generateMSC-educated macrophages (referred to herein as MEMs). In someembodiments, the MSC are LPS-primed MSCs (LPS-MSCs) which have beencultured in the presence of LPS.

MSCs, fibroblasts, and other cells described herein for use in themethods or compositions of the present invention may be derived orisolated from any suitable source. MSCs and fibroblasts may be isolatedfrom tissues including but not limited to bone marrow, lung, cornea,intestines, testis, tendon, adipose, muscle, liver, vertebral,umbilical, and amniotic. In one embodiment, MSCs are isolated from bonemarrow (BM-MSCs). In one embodiment, MSCs are differentiated fromembryonic- or induced pluripotent stem cells.

As used herein, “LPS mesenchymal stem cells (LPS-MSC)” refers to amesenchymal stem cell that has been cultured in the presence of LPS. TheMSCs may be cultured in the presence of LPS for at least about 2 hours.In some embodiments, the MSCs may be cultured in the presence of LPS forat least 12 hours (e.g., at least 10 hours, at least 12 hours, at least15 hours, at least 18 hours, at least 24 hours, or at least 32 hours) inany suitable culture medium known in the art that will support thegrowth and survival of the MSCs. The LPS is present in the culturemedium at a concentration of about 1 ng/ml to about 10 ug/ml.

The LPS used in priming of MSCs as described herein may be from anysuitable source. Suitable sources include, but are not limited to, LPSfrom gram negative bacteria including Escherichia, Salmonella.Neisseria, Hemophilus, Klebsiella, Campylobacter, Bacteroides,Kelicabacter, Pseudomonas, Yersinia, and Shigella. It is known in theart that LPS from various sources have varying degrees of endotoxicactivity. A skilled artisan will recognize that in certain embodimentsit may be advantageous to select a suitable LPS based on the endotoxicactivity thereof.

In some embodiments, LPS may be replaced with another TLR4 ligand.Exemplary TLR4 ligands for use in the methods for priming MSCs describedherein include LPS, VSV glycoprotein G, RSV fusion protein, MMTVenvelope protein, mannan, glucuronoxylomannan,glycosylinositolphospholipids, HSP60, HSP70, fibrinogen, nickel, HMGB1(from Lee, C C et al, Nature Reviews Immunology 12, 168-172 (2012)),12105 (a substituted pyrimido[5,4-b]indole (Goff P H et al. J ofVirology 89, 6 2015)), Glucopyranosyl Lipid Adjuvant (GLA) (Arias, M Aet al. Plos One, PLOS ONE 7(7): e41144. 2012), and synthetic lipid Amimetics (aminoalkyl glucosaminide 4-phosphates, Evans J T et al., J ofExpert Review of Vaccines, 2, 2003).

As used herein, “extraceilular factors” refers collectively toextracellular vesicles, exosomes, micro-vesicles, extracellular matrixcompositions, isolated extracellular matrix components and fragments orderivatives thereof, exosomes purified from an extracellular matrix, andcombinations thereof. Extracellular factors are used in co-culture withCD14⁺ cells to educate macrophages or monocytes in a tissue-specificmanner. As used herein, “extracellular vesicles (EVs)” refers to bothexosomes and micro-vesicles.

As used herein, “exosomes” refer to small lipid vesicles released by avariety of cell types. Exosomes are generated by inward- or reversebudding, resulting in particles that contain cytosol and exposedextracellular domains of certain membrane-associated proteins(Stoorvogel et al., Traffic 3:321-330 (2002)). Methods of preparingexosomes from cells are known in the art. See, for example, Raposo etal., J. Exp. Med. 183:1161 (1996). In one method, exosomes are recoveredfrom conditioned culture medium by centrifugation. Exosomes suited foruse in the methods can be derived fresh or can be previously frozenaliquots kept as a composition, thawed, and added in a single dose orrepeated doses to CD14⁺ cultures to generate educated macrophages. Insome embodiments, exosome preparations may also include micro-vesicles.

Exosomes can have, but are not limited to, a diameter of about 10-300nm. In some embodiments, the exosomes can have, but are not limited to,a diameter between 20-250 nm, 30-200 nm or about 50-150 nm. Exosomes maybe isolated or derived from any cell type that resides in the targettissue of interest which can be isolated and cultured for a period oftime appropriate for the isolation of exosomes.

In one embodiment, the exosomes (LPS-EVs) are derived from MSCs primedwith LPS (LPS-MSCs). In some embodiments, EVs are isolated from the LPSprimed MSC culture by harvesting medium containing the EVs. Multiplecycles of EV isolation may be performed from a single population of MSCsin culture. For example, medium harvested from MSCs after a first LPSpriming can be replaced with fresh media containing LPS for anotherround of LPS priming and EV isolation. In some embodiments, MSCs may beprimed with LPS at a concentration of about 50 ng/ml to about 200 ng/ml(LPS-low). In some embodiments, MSCs may be primed with LPS at aconcentration of about 50 ng/ml, about 100 ng/ml, about 125 ng/ml, about150 ng/ml, about 175 ng/ml, or about 200 ng/ml (LPS-low). In someembodiments, MSCs may be primed with LPS at a concentration of about 800ng/ml to about 1200 ng/ml (LPS-High). In some embodiments, MSCs may beprimed with LPS at a concentration of at least about 800 ng/ml, at leastabout 850 ng/ml, at least about 900 ng/ml, at least about 950 ng/ml atleast about 1000 ng/ml, at least about 1050 ng/ml, at least about 1100ng/ml, at least about 1150 ng/ml, or at least about 1200 ng/ml(LPS-high). In some embodiments MSCs may be primed with LPS at aconcentration of at most about 1200 ng/ml, at most about 1100 ng/ml, atmost about 1050 ng/ml, or at most about 1000 ng/ml. When surface markersare examined by flow cytometry, the LPS-high-EVs are positive for CD105,CD146, CD29, CD44, CD63, CD81, MSCP and CD9.

LPS-exosomes derived from LPS-MSCs (LPS-EVs) are co-cultured with CD14+cells to generate LPS-specific exosome-educated macrophages (referred toherein as LPS-EEMs), which are immunosuppressive, reparative,anti-inflammatory macrophages. LPS-low-EEMs are generated fromLPS-low-EVs and LPS-high-EEMs are generated from LPS-high-EVs, whereinhigh and low indicate the relative concentration of LPS used to cultureMSCs. When comparing by flow cytometry the external surface markers ofLPS-low EEMs to the markers of control macrophages, the LPS-low EEMsshow a significant increase in the intensity of marker expression (MFI)of CD206, PD-L1, and CD16 with a significant decreases in M1 markers inCD86 and HLA-DR. LPS-low EEMs also show a significant increase inpercentage of cells (% cells) expressing CD206, PD-L1, PD-L2, CD16, andCD73 as well as a significant decrease in CD86. Comparing surfacemarkers of control macrophages to LPS-high EEMs, there were significantdecreases in the MFI of CD86 and HLA-DR with significant increases inthe % cells expressing CD73 but decreases in CD86. Gene expressionanalysis of LPS-high-EEMs and LPS-low-EEMs by qPCR shows statisticalincreases in IL-10, IDO, IL-6, VEGF-A, Stat1 and Stat3, TNF-alpha andIL-8, and a statistical decreases in IL-12 as compared to controlmacrophages. Comparing secreted cytokine/chemokine profile to controlmacrophages by multiplex ELISA demonstrated significant increases inLPS-low and/or LPS-high EEMs of the following analytes; EGF, FGF-2,EOTAXIN, TGF-a, G-CSF, FLT-3L, GM-CSF, FRACTALKINE, INFa2, IFNg, GRO,IL-10, MCP-3, IL-12p40, IL-12p70, IL-13, PDGF-BB, IL-15, sCD40L IL-17,IL-1a, IL-1b, IL-2, IL-4, IL-5, IL-7, IL-8, IP-10, MIP-1a, MIP-1b, TNFa,and VEGF. The functions of analytes secreted in high levels by theLPS-EEMs include: growth factors for wound healing (EGF, FGF-2, TGF-a),vascular growth factors (VEGF-A), hematopoietic growth factors (G-CSF,GM-CSF, FLT-3L, IL-7) chemotactic or chemoattractant chemokines(EOTAXIN, FRACTALKINE, GRO, MCP-3, IP-10), anti-inflammatory cytokines(IL-4, IL-10, IL-13) immuno-modulating factors (INFa2, IFNg, IL-17,IL-1a, IL-9, IL-5) and platelet activating factors (PDGF-BB, sCD40L).

LPS-EVs are co-cultured with CD14+ monocytes to generate LPS-specificexosome-educated monocytes (referred to herein as LPS-EEMos), which areimmunosuppressive, reparative, anti-inflammatory monocytes.LPS-low-EEMos are generated from LPS-low-EVs and LPS-high-EEMos aregenerated from LPS-high-EVs, wherein high and low indicate the relativeconcentration of LPS used to culture MSCs. When compared by flowcytometry, the external surface markers of LPS-high-EEMos show asignificant increase in the percentage of cells positive for CD73 andPD-L1 and a significant decrease in expression of CD163, CD16, CD206,PD-L2 and CD86 compared to control monocytes. By flow cytometry, theexternal surface markers of LPS-high-EEMos show a significant increasein the percentage of cells positive for PD-L1 and significant decreasesin CD16, CD206, CD86 and CD73 compared to EEMos generated by co-culturewith EVs derived from MSCs that have not been primed with LPS. Geneexpression studies of the LPS-high-EEMos by qPCR showed statisticalincreases in IL-6, IDO, FGF2, IL-10, and IL-15 compared to both thecontrol monocytes and EEMos. VEGF-A was also statistically higher in theLPS-EEMos compared to control monocytes.

Characteristic surface marker phenotypes and cytokine growth factorprofiles of some embodiments of the educated macrophages describedherein are outlined Example 1.

Treatment

According to the methods of the present invention, educated macrophages,educated monocytes, LPS-EVs or a combination of any two or more of theforegoing of are administered to a subject in need of thereof. Subjectsin need of treatment include those already having or diagnosed with adisease or injury as described herein or those who are at risk ofdeveloping a disease or injury as described herein.

A disease or injury of the present invention may include, but is notlimited to, conditions associated with radiation-induced injury andacute radiation syndrome.

With respect to radiation-induced injury, an amount of ionizingradiation exposure resulting in radiation-induced conditions appropriatefor treatment or prevention according to a method provided herein isgenerally between minimal and maximal tolerance doses. The minimaltolerance dose (T/D_(5/5)) is the dose that when administered to a givenpatient population under a standard set of treatment conditions, resultsin a rate of severe complications of 5% or less within 5 years oftreatment. The maximal tolerance dose (T/D_(50/5)) is the dose that whenadministered to a given patient population under a standard set oftreatment conditions, results in a rate of severe complications of 50%or less within 5 years of treatment. T/D_(5/5) and T/D_(50/5) have beenestablished for many conditions and are well-known (see, e.g., Rubin etal. (Eds) Radiation Biology and Radiation Pathology Syllabus, set RT 1Radiation Oncology, Chicago, American College of Radiology, 1975). Theminimal tolerance dose and maximal tolerance dose have been establishedwith respect to therapeutic radiation treatments but are applicable aswell for determining the range of radiation exposure suitable forcausing the radiation-induced disorders resulting from exposure toradiation from other sources (e.g., occupational or environmentalexposures).

Radiation is quantitated on the basis of the amount of radiationabsorbed by the body, not based on the amount of radiation produced bythe source. A rad (radiation absorbed dose) is 100 ergs of energy pergram of tissue; a gray (Gy) is 100 rad. Radiation dose can be measuredby placing detectors on the body surface or by calculating the dosebased on radiating phantoms that resemble human form and substance.Radiation dose has three components: total absorbed dose, number offractions, and time. Most teletherapy radiation therapy programs arefractionated, being delivered in fractions periodically over time,typically once a day, 5 days a week, in 150-200 cGy fractions, generallyapplied to limited target areas of the body. The total dose delivered inradiation therapy will vary depending on the nature and severity of thecondition being treated. For curative cases, the absorbed dose typicallywill range from 20-80 Gy. For preventative cases, doses are typicallyaround 45-60 Gy and are applied in fractions of about 1.8-2 Gy per day.When used for radiation therapy, ionizing radiation is usually providedover a period of time or until a particular amount of radiation exposurehas been reached by the target area of the subject. Sources of ionizingradiation include electrons, X-rays, gamma rays, and atomic ions.Exposure of a subject to ionizing radiation may be due to a medicalprocedure including, but not limited to, radiation therapy to treatcertain malignant conditions, e.g., lung or breast cancer; medicalprocedures such as diagnostic X-rays; or procedures involvingadministration of nuclear medicines. Exposure to ionizing radiation alsocan result from a nuclear accident or from known or suspectedoccupational or environmental sources, e.g., various consumer productsincluding, but not limited to, tobacco, combustible fuels, smokedetectors, and building materials.

Radiation-induced disorders appropriate for treatment with methods ofthe present invention can result from exposure to ionizing radiation inthe course of radiation therapy. As used herein, the term “radiationtherapy” refers to the medical use of high-energy ionizing radiation toshrink tumors, to control malignant cell growth, or, where appropriate,to treat non-malignant conditions such as thyroid eye disease orpigmented villonodular synovitis. X-rays, gamma rays, and chargedparticles are types of radiation used for radiation therapy. Theradiation may be delivered by a machine outside the body (external-beamradiation therapy, also called teletherapy), or it may come fromencapsulated radioactive material implanted directly into or adjacent totumor tissues in the body near cancer cells (internal radiation therapy,also called brachytherapy). Systemic radiation therapy uses radioactivesubstances, such as radioactive iodine, that travel in the blood and aretargeted in some fashion to the cancer cells. Teletherapy is the mostcommon form of radiation therapy. About half of all cancer patientsreceive some type of radiation therapy sometime during the course oftheir treatment.

Radiation-induced disorders in different tissues and organs generallyfollow a similar course after exposure to ionizing radiation,particularly as a consequence of radiation therapy. Depending on thedose of ionizing radiation to which the subject is exposed, the subjectexperiences an acute response phase that generally occurs days to weeksfollowing exposure to ionizing radiation. The acute response phasetypically involves inflammatory components, and, if low dose, in somepatients, can resolve within a relatively short time or can be fatal.Depending on the dose of ionizing radiation to which the subject isexposed, the acute phase may be followed by a chronic phase, generallybeginning one or more months after exposure. The chronic phase is oftencharacterized by extensive tissue remodeling and fibrosis. Resultspresented herein suggest that effective treatment of the acute responsemay mitigate or attenuate the chronic phase. Cancers or tumors thatoccasionally develop, often many years later, at or near the site ofradiation exposure are not intended to be included among the disorderssuitable for treatment in the method of the present invention.Radiation-induced disorders, particularly those resulting from radiationtherapy, are well known and have been observed in a variety of tissuesand organs. The radiation-induced disorder is not the intended result ofthe radiation therapy but rather is an unintended, and undesirable, sideeffect of the exposure of various organs, tissues and body parts to theionizing radiation used in radiation therapy. The radiation-induceddisorder can be a disorder induced by irradiation of any, or multiple,body parts, organs or tissues of the subject, including but not limitedto bone marrow, lung, heart, bladder, gastrointestinal tract, largeintestine, small intestine, stomach, esophagus, skin, ovaries, testes,urogenital system, kidney, head, neck, pancreas, liver, brain, spinalcord, prostate, vasculature, and muscle. In various aspects theradiation-induced disorder can be, but is not limited to one or more ofbone marrow failure, radiation pneumonitis, radiation enteritis,radiation enteropathy, radiation enterocolitis, radiation dermatitis,radiation-induced erythema, radiation colitis, radiation proctitis,radiation cystitis, radiation nephritis, radiation esophagitis,radiation pericarditis, radiation-induced cardiac effusion, andradiation-induced cardiac fibrosis. All of these disorders arewell-known and readily identifiable by competent medical practitioners.

As used herein, the terms “treat” and “treating” refer to therapeuticmeasures, wherein the object is to slow down or alleviate (lessen) anundesired physiological change or pathological disorder resulting from adisease or injury as described herein. For purposes of this invention,treating the disease, condition, or injury includes, without limitation,alleviating one or more clinical indications, decreasing inflammation,reducing the severity of one or more clinical indications of the diseaseor injury, diminishing the extent of the condition, stabilizing thesubject's disease or injury (i.e., not worsening), delay or slowing,halting, or reversing the disease or injury and bringing about partialor complete remission of the disease or injury. Treating the disease orinjury also includes prolonging survival by days, weeks, months, oryears as compared to prognosis if treated according to standard medicalpractice not incorporating treatment with educated macrophages.

Subjects in need of treatment can include those already having ordiagnosed with a disease or injury as described herein as well as thoseprone to, likely to develop, or suspected of having a disease or injuryas described herein. Pre-treating or preventing a disease or injuryaccording to a method of the present invention includes initiating theadministration of a therapeutic (e.g., human educated macrophages) at atime prior to the appearance or existence of the disease or injury, orprior to the exposure of a subject to factors known to induce thedisease or injury. Pre-treating the disorder is particularly applicableto subjects at risk of having or acquiring the disease injury.

As used herein, the terms “prevent” and “preventing” refer toprophylactic or preventive measures intended to inhibit undesirablephysiological changes or the development of a disorder or conditionresulting in the disease or injury. In exemplary embodiments, preventingthe disease or injury comprises initiating the administration of atherapeutic (e.g., educated macrophages) at a time prior to theappearance or existence of the disease or injury such that the diseaseor injury, or its symptoms, pathological features, consequences, oradverse effects do not occur. In such cases, a method of the inventionfor preventing the disease or injury comprises administering educatedmacrophages to a subject in need thereof prior to exposure of thesubject to factors that influence the development of the disease orinjury.

As used herein, the terms “subject” or “patient” are usedinterchangeably and can encompass any vertebrate including, withoutlimitation, humans, mammals, reptiles, amphibians, and fish. However,advantageously, the subject or patient is a mammal such as a human, or amammal such as a domesticated mammal, e.g., dog, cat, horse, and thelike, or livestock, e.g., cow, sheep, pig, and the like. In exemplaryembodiments, the subject is a human. As used herein, the phrase “in needthereof” indicates the state of the subject, wherein therapeutic orpreventative measures are desirable. Such a state can include, but isnot limited to, subjects having a disease or injury as described hereinor a pathological symptom or feature associated with a disease or injuryas described herein.

In some cases, a method of treating or preventing a disease or injury asdescribed herein comprises administering a pharmaceutical compositioncomprising a therapeutically effective amount of educated macrophages,educated monocytes, LPS-EVs, or a combination thereof as a therapeuticagent (i.e., for therapeutic applications). As used herein, the term“pharmaceutical composition” refers to a chemical or biologicalcomposition suitable for administration to a mammal. Examples ofcompositions appropriate for such therapeutic applications includepreparations for parenteral, subcutaneous, transdermal, intradermal,intramuscular, intracoronarial, intramyocardial, intraperitoneal,intravenous or intraarterial (e.g., injectable), or intratrachealadministration, such as sterile suspensions, emulsions, and aerosols.Intratracheal administration can involve contacting or exposing lungtissue, e.g., pulmonary alveoli, to a pharmaceutical compositioncomprising a therapeutically effective amount of educated macrophages oreducated monocytes alone or in combination with LPS-EVs. In some cases,pharmaceutical compositions appropriate for therapeutic applications maybe in admixture with one or more pharmaceutically acceptable excipients,diluents, or carriers such as sterile water, physiological saline,glucose or the like. For example, educated macrophages described hereincan be administered to a subject as a pharmaceutical compositioncomprising a carrier solution.

Formulations may be designed or intended for oral, rectal, nasal,topical or transmucosal (including buccal, sublingual, ocular, vaginaland rectal) and parenteral (including subcutaneous, intramuscular,intravenous, intraarterial, intradermal, intraperitoneal, intrathecal,intraocular and epidural) administration. In general, aqueous andnon-aqueous liquid or cream formulations are delivered by a parenteral,oral or topical route. In other embodiments, the compositions may bepresent as an aqueous or a non-aqueous liquid formulation or a solidformulation suitable for administration by any route, e.g., oral,topical, buccal, sublingual, parenteral, aerosol, a depot such as asubcutaneous depot or an intraperitoneal or intramuscular depot. In somecases, pharmaceutical compositions are lyophilized. In other cases,pharmaceutical compositions as provided herein contain auxiliarysubstances such as wetting or emulsifying agents, pH buffering agents,gelling or viscosity enhancing additives, preservatives, flavoringagents, colors, and the like, depending upon the route of administrationand the preparation desired. The pharmaceutical compositions may beformulated according to conventional pharmaceutical practice (see, e.g.,Remington: The Science and Practice of Pharmacy, 20th edition, 2000, ed.A. R. Gennaro, Lippincott Williams & Wilkins, Philadelphia, andEncyclopedia of Pharmaceutical Technology, eds. J. Swarbrick and J. C.Boylan, 1988-1999, Marcel Dekker, New York).

The preferred route may vary with, for example, the subject'spathological condition or weight or the subject's response to therapy orthat is appropriate to the circumstances. The formulations can also beadministered by two or more routes, where the delivery methods areessentially simultaneous or they may be essentially sequential withlittle or no temporal overlap in the times at which the composition isadministered to the subject.

Suitable regimes for initial administration and further doses or forsequential administrations also are variable, may include an initialadministration followed by subsequent administrations, but nonetheless,may be ascertained by the skilled artisan from this disclosure, thedocuments cited herein, and the knowledge in the art.

In some cases, educated macrophages, educated monocytes, LPS-EVs orcombinations thereof may be optionally administered in combination withone or more additional active agents, including exosomes ormicrovesicles. Such active agents include anti-inflammatory,anti-cytokine, analgesic, antipyretic, antibiotic, immunosuppressiveagents and antiviral agents, as well as growth factors and agonists,antagonists, and modulators of immunoregulatory agents (e.g., TNF-α,1L-2, IL-4, IL-6, IL-10, IL-12, IL-13, IL-18, IFN-α, IFN-γ, BAFF,CXCL13, IP-10, VEGF, EPO, EGF, HRG, Hepatocyte Growth Factor (HGF),Hepcidin, including antibodies reactive against any of the foregoing,and antibodies reactive against any of their receptors). Any suitablecombination of such active agents is also contemplated. Whenadministered in combination with one or more active agents, educatedmacrophages can be administered either simultaneously or sequentiallywith other active agents. For example, victims of acute radiationsyndrome may simultaneously receive educated macrophages and a growthfactor (such as G-CSF or PEG-G-CSF), a cytokine (such as IL-3, IL-11,IL-12), a population of cells (such as lymphoid or myeloid progenitors),or a small molecule radio-protective agent (such as amafostine orgenistein) for a length of time or according to a dosage regimensufficient to support recovery and to treat, alleviate, or lessen theseverity of the radiation injury. In some embodiments, educatedmacrophages, educated monocytes, or LPS-EVs of the present invention mayalso be administered to a patient simultaneously with or prior toreceiving a radiation treatment, such as a treatment for cancer. In someembodiments, CD14+ monocytes or macrophages are administeredsimultaneously with LPS-EVs to a patient.

In some embodiments, educated macrophages, educated monocytes, LPS-EVsor a combination thereof are administered to a subject in need thereofusing an infusion, topical application, surgical transplantation, orimplantation. In an exemplary embodiments, administration is systemic.In such cases, educated macrophages, educated monocytes. LPS-EVs or acombination thereof can be provided to a subject in need thereof in apharmaceutical composition adapted for intravenous administration tosubjects. Typically, compositions for intravenous administration aresolutions in sterile isotonic aqueous buffer. The use of such buffersand diluents is well known in the art. Where necessary, the compositionmay also include a local anesthetic to ameliorate any pain at the siteof the injection. Generally, the ingredients are supplied eitherseparately or mixed together in unit dosage form, for example, as acryopreserved concentrate in a hermetically sealed container such as anampoule indicating the quantity of active agent. Where the compositionis to be administered by infusion, it can be dispensed with an infusionbottle containing sterile pharmaceutical grade water or saline. Wherethe composition is administered by injection, an ampoule of sterilewater for injection or saline can be provided so that the ingredientsmay be mixed prior to administration. In some cases, compositionscomprising human educated macrophages, educated monocytes, LPS-EVs orcombinations thereof are cryopreserved prior to administration.

Therapeutically effective amounts of educated macrophages, educatedmonocytes, LPS-EVs or combinations thereof are administered to a subjectin need thereof. An effective dose or amount is an amount sufficient toeffect a beneficial or desired clinical result. With regard to methodsof the present invention, the effective dose or amount, which can beadministered in one or more administrations, is the amount of humaneducated macrophages, educated monocytes, or LPS-EVs sufficient toelicit a therapeutic effect in a subject to whom the cells areadministered. In some cases, an effective dose of educated macrophagesor educated monocytes is about 1×10⁵ cells/kilogram to about 10×10⁹cells/kilogram of body weight of the recipient. In some cases, aneffective dose of LPS-EVs is about 1×10⁵ extracellular vesicles/kilogramto about 10×10¹⁰ extracellular vesicles/kilogram body weight of therecipient. Effective amounts will be affected by various factors thatmodify the action of the cells upon administration and the subject'sbiological response to the cells, e.g., severity of radiation injury,type of damaged tissue, the patient's age, sex, and diet, time ofadministration, and other clinical factors.

Therapeutically effective amounts for administration to a human subjectcan be determined in animal tests and any art-accepted methods forscaling an amount determined to be effective for an animal for humanadministration. For example, an amount can be initially measured to beeffective in an animal model (e.g., to achieve a beneficial or desiredclinical result). The amount obtained from the animal model can be usedin formulating an effective amount for humans by using conversionfactors known in the art. The effective amount obtained in one animalmodel can also be converted for another animal by using suitableconversion factors such as, for example, body surface area factors.

It is to be understood that, for any particular subject, specific dosageregimes should be adjusted over time according to the individual needand the professional judgment of the person administering or supervisingthe administration of the educated macrophages, educated monocytes, orLPS-EVs. For example, an educated macrophage dosage for a particularsubject with radiation injury can be increased if the lower dose doesnot elicit a detectable or sufficient improvement in one or moresymptoms of radiation injury. Conversely, the dosage can be decreased ifthe radiation injury is treated or eliminated.

In some cases, therapeutically effective amounts of educatedmacrophages, educated monocytes, LPS-EVs or combinations thereof can bedetermined by, for example, measuring the effects of a therapeutic in asubject by incrementally increasing the dosage until the desiredsymptomatic relief level is achieved. A continuing or repeated doseregimen can also be used to achieve or maintain the desired result. Anyother techniques known in the art can be used as well in determining theeffective amount range. Of course, the specific effective amount willvary with such factors as the particular disease state being treated,the physical condition of the subject, the type of animal being treated,the duration of the treatment, and the nature of any concurrent therapy.

Following administration of educated macrophages, educated monocytes,LPS-EVs or combinations thereof to an individual subject afflicted by,prone to, or likely to develop a disease or injury described herein, aclinical symptom or feature associated with the disease or injury isobserved and assessed for a positive or negative change. For example,for methods of radiation injury in a subject, positive or negativechanges in the subject's infection, bleeding or anemia during orfollowing treatment may be determined by any measure known to those ofskill in the art including, without limitation, blood counts.

In any of the methods of the present invention, the donor and therecipient of the educated macrophages, educated monocytes or LPS-EVs canbe a single individual, autologous, or different individuals, forexample, allogeneic or xenogeneic individuals. Stromal cells and CD14⁺cells for use in the present invention do not need to be from the samedonor, patient or source. As used herein, the term “allogeneic” refersto something that is genetically different although belonging to orobtained from the same species (e.g., allogeneic tissue grafts or organtransplants). “Xenogeneic” means the cells could be derived from adifferent species. In one embodiment, CD14+ cells can be collected frompatients and educated to be given fresh to a person following orconcurrently with radiation treatment such as a cancer treatment. Insome embodiments, any allogeneic donor may act as a universal thirdparty donor of CD14+ cells.

The present invention has been described in terms of one or morepreferred embodiments, and it should be appreciated that manyequivalents, alternatives, variations, and modifications, aside fromthose expressly stated, are possible and within the scope of theinvention.

Example 1

The embodiment described here demonstrates the use of LPS-specificeducated macrophages (LPS-EEMs) and LPS-specific monocytes (LPS-EEMos),generated from the co-culture with extracellular vesicles derived fromLPS-primed MSCs, for the treatment of radiation induced injury. Theembodiment described here also demonstrates the use of EVs from MSCswith and without LPS priming for the treatment of radiation inducedinjury.

Materials and Methods

Cell culture—Monocytes were isolated from human peripheral blood usingmagnetic bead separation methods according to manufacturers' protocols.Briefly, peripheral blood mononuclear cells were collected from theblood after mobilization from healthy donors by density gradientseparation using Ficoll-Paque Plus (endotoxin tested) (GE HealthcareBio-Sciences, Piscataway, N.J., USA) using an IRB-approved protocol. Ifperipheral blood has undergone apheresis designed to concentrate whitecells and exclude red blood cells the density gradient separation stepmay be skipped, in which case the cells are diluted in buffer such asPBS, centrifuged at 300-1000×g and the pellet resuspended in ACK lysisbuffer. Red blood cells were lysed by incubating cells in ACK lysisbuffer for 3-5 minutes and mononuclear cells were washed withphosphate-buffered saline (PBS) (Hyclone, Logan, Utah, USA). To reduceplatelet contamination, cell suspensions were centrifuged at 300-700 rpmfor 10 minutes and cell pellets were re-suspended in Miltenyi separationbuffer with anti-human CD14 microbeads as directed by the manufacturer(Miltenyi Biotec, Auburn, Calif., USA) and incubated for 15 minutes at4° C. After washing to remove unbound antibody, cell separation was doneusing an autoMACS Pro Separator (Miltenyi Biotec). Purity of isolatedCD14⁺ cells was >95% when checked by flow cytometry. Purified CD14⁺monocytes were either plated into six-well cell culture plates at aconcentration of 0.5-1×10⁶ per well for characterization studies or 10⁷per T75 cm² filter cap cell culture flask for animal studies (GreinerBio-One, Monroe, N.C., USA) in Iscove's modified Dulbecco's media(Gibco, Life Technologies, Grand Island, N.Y.) supplemented with 10%human serum blood type AB (Mediatech, Herndon, Va., USA or ValleyBiomedical Inc, Winchester, Va., USA), 1× nonessential amino acids(Lonza, Walkersville, Md., USA), 4 mM L-glutamine (Invitrogen, Carlsbad,Calif., USA), 1 mM sodium pyruvate (Mediatech), and 4 ug/mL recombinanthuman insulin (Invitrogen). Cells were cultured for 7 days at 37° C.with 5% CO₂, without cytokines, to allow differentiation to macrophages.Attached cells were harvested using Accumax dissociation media(Innovative Cell Technologies, Inc, San Diego, Calif.).

MSCs were isolated from filters left over after bone marrow (BM) harvestfrom normal healthy donors using an IRB-approved protocol. Briefly, BMcells trapped in the filter were recovered by rinsing the filter withPBS and mononuclear cells were separated using Ficoll-Hypaque 1.073 (GEHealthcare Bio-Sciences). Red blood cells were lysed with 3-minuteincubation in ACK lysis buffer (Lonza, Walkersville, Md., USA) andmononuclear cells were suspended in α-minimum essential mediumsupplemented with 10% fetal bovine serum (US origin, uncharacterized;Hyclone, Logan, Utah, USA), 1× nonessential amino acids, and 4 mML-glutamine. Cells were cultivated in 75-cm² filter cap cell cultureflasks. Attached cells (passage 0) were harvested using TrypLE™ celldissociation enzyme (invitrogen) and then re-plated into new flasks asdescribed previously²⁰. Passage 4-6 cells were used for characterizationstudies and used for isolation of extracellular vesicles (EVs). Theidentity of the MSCs was confirmed by flow cytometry, and theirimmune-modulatory properties on T-cell proliferation were confirmed byan immunopotency assay.^(39,40)

Isolation and characterization of EVs from cells—Cells (either MSCs ormacrophages) in 75-cm² filter cap cell culture flasks were washed oncewith PBS, and the medium was replaced with StemPro® MSC serum-free media(SFM) CTS (A103332-01, Gibco Life Technologies). Cells were incubatedfor 18-24 hours and the conditioned culture media (CM) was collected. Toprime MSCs with a TLR4 ligand, lipopolysaccharide (LPS) at two differentconcentrations was co-cultivated with MSCs to produce LPS exosomes. SFMwas supplemented with either 100 ng/ml (LPS-low) or 1.0 ug ml (LPS-high)E. coli LPS O111:B4 (L4391 Sigma, St Louis, Mo., USA). EVs were isolatedfrom un-primed MSCs (MSC-EV), LPS-primed MSCs (LPS-low or high EVs), orun-primed macrophages (macrophage-EV) by a 2-step centrifugation processas described.⁴¹ Briefly, the CM was centrifuged using an Allegra® X-15Rcentrifuge (Beckman Coulter, Indianapolis Ind., USA) at 2000×g at 4° C.for 20 minutes to remove any detached cells, apoptotic bodies and celldebris. Clarified supernatant CM was centrifuged in an Optima™ L-80XPUltracentrifuge (Beckman Coulter) at 100,000×g avg at 4° C. for 2 hourswith a SW 28 rotor to pellet exosomes. The supernatant was carefullyremoved, and EV-containing pellets were re-suspended in PBS and pooled.We typically suspended the EV pellet at 100 ul PBS/10 ml of CM whichgave EV particle concentrations of about 10¹⁰ particles/ml (see Table1). To visualize the EVs by transmission electron microscopy (TEM), there-suspended EVs were layered on a 30% sucrose cushion andre-centrifuged at 100,000 g_(avg) at 4° C. for 2 hours. The upperportion of the cushion was collected and re-centrifuged. The pellet wasresuspended in a small volume of PBS, whole mounted on Formvar EM gridsand stained with uranyl acetate as described.⁴¹

EVs were characterized for protein and RNA concentration using aNanoDrop spectrophotometer (Thermo-Fisher, Waltham, Mass., USA). Meanand mode particle diameter and concentration (EV particles/mi) wereassessed using an IZON qNano Nanoparticle Characterization instrument(Cambridge. Mass., USA) or a Nanosight NS300 (Malvern, UK). Thisanalysis coupled with TEM indicated that the vast majority of EVpreparation consisted of exosome-sized vesicles. Therefore, the EVs werealso identified as exosomes and used synonymously.

Characterization of EV (exosome) surface marker profile by MACsplex—Thesurface marker profile of EVs from two MSC isolates of both unstimulatedMSC-EVs and LPS-high EVs were determined by flow cytometry using theMACSPlex Exosome Kit (Miltenyi Biotec). This kit allows the detection of37 exosomal surface markers and two isotype markers that served asisotype controls (D1c, CD2, CD3, CD4, CD8, CD9, CD1c, CD14, CD19, CD20,CD24, CD25, CD29, CD31, CD40, CD41b, CD42a, CD44, CD45, CD49e, CD56,CD62P, CD63, CD69, CD81, CD86, CD105, CD133/1, CD142, CD146, CD209,CD326, HLA-ABC, HLA-DRDPDQ, MCSP, ROR1, SSEA-4, REA control, mlgG1control). This assay was performed according to manufacturer's protocol.In brief, capture beads coupled with antibodies to the exosome surfacemarkers were mixed with equal volumes of purified MSC exosomes andgently rotated in the dark at 4° C. overnight. The bead-exosomecomplexes were washed and then incubated for 1 hour with detection beadmixture consisting of pan-exosome markers CD9, CD63 and CD81 labeledwith FITC, PE or APC. The beads were then washed and resuspended in 150uls of MACSPIex buffer for analysis. Prior to experimentation, thesystem was calibrated and background settings were adjusted to unlabeledbeads. The auto-sampler used 100 uls from each sample to collect beadsand automated gating strategies were used to identify bead populationsfor each analyte. Batch analysis quantified median intensities for eachbead population and analyte surface expression was calculated for eachsample. Miltenyi MACSQuant Analyzer 10 for sample acquisition andMACSQuantify Software was used for data analysis. Median fluorescentvalues from exosomes isolated from different MSC isolates were averagedand values of 1.0 or more were considered significant compared tobackground.

Education of CD14⁺ cells by co-culture with EVs or MSCs—For education ofCD14⁺ macrophages, frozen stocks of CD14⁺ monocytes were thawed, thenplaced in complete macrophage media and allowed to differentiate toadherent macrophages for 5-7 days. These macrophages were thensupplemented with fresh media and educated for an additional 3 days withEVs. For education of CD14⁺ monocytes, the frozen stocks of CD14⁺monocytes were thawed, then placed in complete macrophage media andtreated within 1 hour with EVs and educated for 18-24 hours. Thetypically EV stock concentration was 10¹⁰ particles/ml in PBS, based onEV particle concentration/ml determined using the IZON qNanoNanoparticle Characterization instrument (Cambridge, Mass., USA). Theamount of the EV preparation used for education was based ondose-response studies using EVs coupled with flow-cytometry to determinechanges in surface marker expression. For education, 40-60 ul of EVswere used for 6 well plates (2 mls media) or 250-300 ul of EVs in 75-cm²filter cap cell culture flasks (10 mls of media). EVs were isolated fromeither unstimulated MSCs (MSC-EVs), MSCs primed with LPS, (LPS-low-EVs,LPS-high-EVs) or from Day 10 macrophages to generate macrophage-EVs.Macrophage-EV preparations served as non-MSC control EVs. Educatedmacrophages generated by co-culture with EVs from various sources(MSC-EVs, LPS-low-EVs, LPS-high-EVs or macrophage-EVs) were designatedas EEMs, LPS-low EEMs, LPS-high EEMs and macrophage-EEMs, respectively.Educated monocytes generated by co-culture with EVs from various sources(MSC-EVs or LPS-high-EVs) were designated as EEMos and LPS-high-EEMos,respectively. Direct co-culture of CD14⁺ macrophages with MSCs generatedMSC-educated macrophages (MEMs). Day 7 macrophages were supplementedwith fresh media and incubated with human BM-derived MSCs at anapproximate ratio of 10:1 of macrophages: MSCs, and incubated for 3 daysas previously described to generate MEMs.²⁰

M1 stimulation of macrophages—In contrast to treating Day 7 macrophageswith EVs of MSCs, Day 7 macrophages (M0, naïve macrophages) weredirectly stimulated with pro-inflammatory agents to produce an M1phenotype and serve as a comparison to the macrophages educated by EVsor MSCs. Fresh medium was added supplemented with 320 nM phorbol12-myristate 13-acetate (PMA) for 6 hours followed by the addition of 20ng/ml Interferon-gamma and 100 ng/ml LPS and incubated for at least 18hours.⁴² These macrophages directly stimulated with M1 factors (M1stimulated) were used in gene expression analysis studies for comparisonto control macrophages, EEMs, LPS-low-EEMs and LPS-high-EEMs. The M1stimulated macrophages were also tested in the phagocytosis assay andthe results compared to control macrophages, MEMs, EEMs, LPS-low-EEMsand LPS-high-EEMs.

Cells were harvested by removing media, washing with phosphate-bufferedsaline (PBS, Hyclone) then using Accumax cell dissociation enzyme(Innovative Cell Technologies, Inc, (San Diego, Calif., USA) to detachthe cells from the flask followed by the use of a cell scraper. Aportion of the cells was analyzed by flow cytometry and the remainderwas used for animal studies.

Flow Cytometry—Table 3 and FIGS. 3A-3B show macrophage flow cytometrydata. FIG. 4 shows monocyte flow cytometry data. Macrophages (controls(MO), MEMs, EEMs, LPS-high-EEMs, and LPS-low-EEMs) at day 10 of cultureand monocytes (controls (Mo), EEMos, and LPS-high-EEMos) at day 2 ofculture were collected, counted and incubated with Fe block (BDPharmingen, San Jose, Calif., USA, cat #: 564220) and stained at 4° C.for 20-30 minutes with anti-human antibodies. All antibodies werepurchased from BioLegend (San Diego, Calif.) except BV510-CD73 (AD2, cat#563198) from BD Pharmingen. Panels included: MSCs, PE-Cy7-CD90 (5E10,cat #328123), macrophages, PerCP/Cy5.5-CD14 (HCD14, cat #325622),BV421-CD16 (3G8, cat #302038), M2 markers, FITC-CD163 (GHI/61, cat#333618), FITC-CD39 (A1, cat #328206), PE-CD206 (15-2, cat #321106),APC-PD-L1 (29E.2A3, cat #329708), APC-PD-L2 (24F.10C12, cat #329608), M1markers, BV510-CD86 (IT2.2, cat #305432) and Pacific Blue-HLA-DR/MHC 11(L234, cat #307633). For MEM marker analysis, macrophages (CD14CD90-)were selectively gated using CD14 and CD90 to exclude the MSCs(CD14-CD90-). Flow cytometry data were acquired on a MACSQuant analyzer10 (Miltenyi Biotec). Mqd files were converted to fcs files using TheMACSQuantify™ Software. Listmode data were analyzed using FlowJo™software (TreeStar).

Gene expression analysis—RNA was isolated from cells using RNeasy™ microkit (Qiagen, Valencia, Calif., USA), and the quality of isolated RNA waschecked using Nanodrop 1000 (Fisher Scientific, Pittsburgh, Pa., USA).RNA was converted to cDNA using Quantitect reverse transcription kit(Qiagen). Quantitative polymerase chain reaction was performed usingPower SYBR green master mix (Applied Biosystems, Foster City, Calif.,USA) on StepOne Plus instrument (Applied Biosystems) using standardprotocols. Verified primers for IL-10, indoleamine 2,3-dioxygenase(110), IL1B, IL-6, IL-8, IL-10, IL-12, IL-23, Serpine-1, TGF-B, TNF-α,Stat 1, Stat 3 and VEGF-A were purchased from Qiagen. The thresholdcycle (Ct) value for each gene was normalized by the average Ct usingthe GAPDH housekeeping gene and using this normalization the expressionvalues of the control macrophages or control monocytes were set at 1.0.

Phagocytic assay—Day 7 macrophages were either untreated, stimulatedwith M1 factors, or educated by co-cultivation with MSCs, or usingMSC-EVs to generate EEM, LPS-low EEM or LPS-high EEM for 3 days asdescribed above. The phagocytic assays were performed on Day 10macrophages using the pHrodo Green E. coli Bioparticle conjugate system(cat #P35366, Invitrogen) according to manufacturer recommendations. Thefluorescence of the pHrodo Green is activated within the phagosome ofthe cell as the pH decreases and reduces the detection of non-phagocyticbinding. The pHrodo Green E. coli Bioparticle conjugate wasreconstituted in PBS, diluted in media, and incubated for 1 hour at 37C. Cells were washed with cold PBS three times to reduce non-specificattachment collected by cell scraping. Collected cells were treated withFc Receptor blocker for 10 minutes and macrophages stained withCD14-PerCP 5.5 for 20 minutes at 4° C. CD14 positive/pHrodo Greenpositive cells were detected on the MACSQuant analyzer 10 (MiltenyiBiotec) and analyzed using FlowJo™ software (TreeStar).

Multiplex Cytokine ELISA assay—Day 7 macrophages (10⁶/well) grown in6-well plates were either untreated (control), stimulated with M1factors or educated for 3 days in culture media with MSC-EVs to generateEEMs, LPS-low EEMs, and LPS-high EEMs as described in above. The cellswere washed with PBS, medium was replaced and after 24 hours cells wererecovered, centrifuged at 300×g for 10 minutes to remove cell debris andassayed for cytokines and other factors using a Milliplex MAPcytokine/chemokine multiplex magnetic bead panel (HCYTOMAG-60K,Millipore, Burlington Mass.). Twenty-five ul of culture media from threesets of macrophage isolates, each set performed in duplicate wells wereassayed for secreted products as directed by the manufacturer anddetected on a Luminex xMAP platform. The analytes assayed were EGF,FGF-2, EOTAXIN, TGF-a, G-CSF, FLT-3L, GM-CSF, FRACTALKINE, INFa2, IFNg,GRO, IL-10, MCP-3, IL-12p40, MDC, IL-12p70, IL-13, PDGF-BB, IL-15,sCD40L IL-17, IL-1ra, IL-1a, IL-9, IL-1b, IL-2, IL-4, IL-5, IL-6, IL-7,IL-8, IP-10, MCP-1, MIP-1a, MIP-1b, RANTES, TNFa, TNFb and VEGF.

Immunopotency Assay. The immunopotency assay (IPA) was used to determineif the co-culture of T-cells with either MSCs, macrophages, EEMs orLPS-EEMs affects proliferation of CD4+T-helper and/or CD8+T-cytotoxiccells after stimulation with anti-CD3 and anti-CD28. Peripheral bloodmononuclear cells (PBMCs), the source of the CD4+ and CD8+ cells wereisolated from leukapheresis products collected from different normalhealthy donors and were purchased from SeraCare Life Sciences (Milford,Mass.). After Ficoll separation, between 4 and 5×10⁹ PBMCs wererecovered then cryopreserved at 1×10⁷ cells/vial using 90% FBS (AtlantaBiologics, Atlanta, Ga.) and 10% DMSO (Sigma-Aldrich, St. Louis, Mo.)and stored in LN2 before use in this assay. The test cells used inco-cultivation were isolated as described above and consisted of either:MSCs, macrophages, EEMs or LPS-low EEMs. They were all prepared in IPAmedium consisting of RPM1-1640 containing 10% heat inactivated FBS, 1×non-essential amino acids (NEAA)(Mediatech, Inc., Manassas, Va.), 1×Glutamine (Mediatech, Inc.), 1×Na Pyruvate (Sigma-Aldrich), and 1×HEPESbuffer (Sigma-Aldrich, St. Louis, Mo.). This assay was performed in a 48well tissue culture plate with a total IPA medium volume of 400 ul perwell. Preparation of test cells for this assay included wash, andre-suspension at 4×10⁶/mL.

For a 1:1 (PBMC:test cell) ratio, 4-10 MSCs (100 ul) were plated andthen titrated further to 2×10⁴ to achieve a 1:0.05 (PBMC:test cell)ratio. The stimulated PBMC to test cell ratios that were evaluated inthis assay include 1:1, 1:0.5, 1:0.1, and 1:0.05. The volume of testcells was held constant at 200 ul/well using IPA medium. After plating,the test cells were allowed to settle and adhere to the plastic for 2hours at 37° C. To measure proliferation, PBMCs were labeled withcarboxyfluorescein succinate-ester (CFSE) at a final concentration of 1uM for 10 minutes, at 37° C. in the dark, mixing at the 5 minute timepoint to ensure homogeneous labeling. An equal volume of cold FBS wasadded for 1 minute to stop the CFSE labeling reaction. PBMCs were thenwashed twice with IPA medium as defined above before reconstitution at4×10⁴/mL. One hundred microliters of CFSE-labeled PBMCs was added toeach well containing the various ratios of test cells. Anti-CD3 andanti-CD28 antibodies (clones UCHT1 and 37407, respectively) (R&DSystems, Inc., Minneapolis, Minn.) also prepared in the IPA medium wereused to stimulate the proliferation of CD4+ helper T-cells and CD8+cytotoxic T-cells. A 100 ul mixture of 4× concentrated anti-huCD3 andanti-huCD28 antibodies (10 ug/mL and 2 ug/mL, respectively) was added toeach well except for the 1:0.05 (PBMC:test cell) non-stimulated controlwhich received 100 ul of IPA medium instead (negative control). Cellswere cultured for 4 days at 37° C. before the CD4+ T cells were analyzedfor proliferation using standard flow cytometry methodology. Anti-huCD4APC or anti-huCD8 APC (R&D Systems, Inc.) was used to gate the CD4+T andCD8+ T cells. All IPA analyses were performed using an Accuri C6 flowcytometer (BD Biosciences, Inc., San Jose, Calif.) and the associated C6Plus software was used for the CFSE analysis.

Mice—Male and Female NOD.Cg-Prkdc^(scid)Il2rg^(tmlWjl)/SzJ (NSG) micewere purchased from The Jackson Laboratory (Bar Harbor, Me.) and used at8-16 weeks of age. All animals were bred and housed in a pathogen-freefacility throughout the study. The Animal Care and Use Committee at theUniversity of Wisconsin approved all experimental protocols.

In vivo lethal radiation injury model—On day 0, approximately equalnumbers of NSG male and female mice received 4 Gy lethal total bodyirradiation using an X-RAD 320 X-ray irradiator (Precision X-Ray, NorthBranford, Conn., USA) to induce consistent lethality within a 2 weektime frame. Four hours after radiation injury, mice were treatedintravenously in the tail vein with either 100 ul of PBS (control),1×10⁶ human macrophages, 1×10⁶ human bone marrow-derived MSCs (passage4-6), 1×10⁶ EEMs, 1×10⁶ LPS-low-EEMs, 1×10⁶ LPS-high-EEMs, 1×10⁷monocytes (Mo), or 1×10⁷ LPS-high-EEMos. For EV treatment studies, micewere treated immediately post radiation challenge with varying dosages(2-5×10¹⁰ of MSC-EVs or LPS-EVs. The mice were monitored at least 3times a week for survival and weight change. Clinical scores were alsodetermined based on a modified clinical scoring system for GVHD.Cumulative scores of percent weight loss, posture, activity, and furtexture (scored from 0-2 for each criteria), were recorded as previouslydescribed⁴³. On day 4 and day 32 post-challenge, blood from survivingmice was collected in a microtainer tube K2 EDTA (Becton Dickenson,Franklin Lakes, N.J.) or equivalent from a tail vein nick and thehematology of the whole blood was assayed on a Hemavet™ 950FS analyzer(Drew Scientific Inc., Miami Lakes, Fla., USA).

Diagnostic necropsy and histologic preparation—Gross necropsy of organsystems consisting determination of both organ weight and organ weightas function of percent body weight (% BW) as well as the externalexamination of the integument, cardiovascular, respiratory, digestive,lympho-hematopoietic, uro-genital, endocrine, central nervous system andmusculoskeletal. Gross necropsy was performed on the following groups:Un-irradiated mice, mice post radiation challenge on day 9 (PBS treatedand LPS-high EEM treated), day 31 (LPS-high EEM treated), and day 52-53(moribund LPS-high EEM treated). At strategic time points post-radiationchallenge, spleens were harvested, weighed and from either moribundcontrol mice or treated mice and compared to the weight of normalun-irradiated healthy controls. Histology focused on the preparations ofslides from sections of spleen and bone marrow from the femur, vertebraeand ileum. Tissues were fixed in 10% neutral buffered formalin andprocessed on a Sakura Tissue-Tek VIP 6 processor and embedded on aSakura Tissue-Tek TEC embedding station. Slides were cut on a Leitz 2235microtome at 5-6 microns and stained with H&E using a Sakura Tissue-TekDRS automatic stainer and manually cover slipped. Tissues werevisualized using a Nikon Eclipse 50 I microscope at multiplemagnifications using Nikon objectives; 4×/0.2—Plan Apo, 10×/0.45 PlanApo, 20×/0.75 Plan Apo, 40×, 0.95 Plan Apo. Photographs were taken usinga SPOT model 10.2 camera aided with SPOT acquisition software for MAC5.2.

Statistical analysis—Statistics were performed using GraphPad Prismversion 7.0 (GraphPad Software, San Diego, Calif.). Data were reportedas mean f SEM. For analysis of three or more groups, the analysis ofvariance (ANOVA) test was performed with the Kruskal-Wallis and Dunn'smultiple comparisons post-test. Principal component analysis and t-testscomparing gene expression between groups were performed on MicrosoftExcel. Multiple hypotheses testing correction was done using theBenjamini Hochberg false discovery rate (FDR) procedure. A p-value lessthan 0.05 was considered statistically significant.

Results

Characterization of the extracellular vesicles(EVs)—Electron-microscopic observation (FIG. 1a,b ) indicate EVs fromthe MSCs have the typical appearance of an EV; circular shape withconvex center and the majority of EV preparation consisted ofexosome-sized vesicles (<200 nm). EVs are composed of exosomes,generally 50-250 nm in size, and larger micro-vesicles typically greaterthan 500 nm. Quantifying the EVs using either a resistive pulse sensinginstrument (qNano Nanoparticle instrument, FIG. 1c ) or a visualnanoparticle tracking analysis (Nanosight NS300, FIG. 1d ) yieldedsimilar profiles in terms of mean particle sizes, range and particledensity. The analysis of EVs preps from multiple MSC isolates using theqNano instrument also indicated that the mean particles size (139nm+/−15) and mode particle size (97 nm+/−9) were consistent withexosome-sized vesicles (Table 1). There were slight differences detectedin mean and mode sizes between MSC isolates ranging from 84 to 181 nm,although mean particle concentrations were very similar at 1.5×10¹¹particles/ml and ranged from 0.76 to 2.0×10¹¹ particles/ml. Unlike aprevious report³⁷, we did not detect a significant increase in theparticle concentration using the qNano instrument after priming of MSCswith LPS at either low or high dosages. Macrophages also produced EVsthat were mostly in the exosomes size range but interestingly secretedabout 10-fold more EVs than MScs based on cell number.

Table 1: This table characterizes the size and concentration of theexosomes isolated from different sources of cells (BM-MSC, macrophages)using the qNano Nanoparticle instrument. The size (mean/mode) andparticle concentrations (/ml) overall are very consistent between batch(culture round, first, third), cell type, or whether the MSCs wereprimed with high or low concentrations of LPS. EVs isolated and analyzedusing qNano Nanoparticle instrument from 4 different BM-MSCs isolates,(15PH05, 15PH06, 15PH07 and 15PH09), primed with LPS-high or low, atdifferent passages (P3-P6), or from different rounds of CM harvest(first or third) all generated preparations with similar yields ofparticle concentration/ml. In addition, the particle number producedbased on cell number (10⁶ cells) was also similar. However, macrophages,which are cultivated at about 10-fold lower cell densities were found toproduce more exosomes based on cell concentration.

Mean Mode particle particle Approx Particle Culture size size Particleconcentration/10⁶ Cell type Isolate round (nm) (nm) concentration/mlcells BM MSC 15PH05 first 92 64 7.6 × 10 ¹⁰ 8.4 × 10 ⁹  P3 BM MSC 15PH06first 162 108 1.6 × 10 ¹¹ 1.7 × 10 ¹⁰ P4 BM MSC 15PH07 first 85 84 1.8 ×10 ¹¹ 2.0 × 10 ¹⁰ P4 BM MSC 15PH09 first 86 61 2.0 × 10 ¹¹ 2.2 × 10 ¹⁰P3 BM MSC 15PH05 first 169 114 1.3 × 10 ¹¹ 1.4 × 10 ¹⁰ P4 BM MSC 15PH05third 175 114 1.4 × 10 ¹¹ 1.5 × 10 ¹⁰ P4 BM MSC 15PH05 third 181 131 1.2× 10 ¹¹ 1.3 × 10 ¹⁰ P4 (LPS- low) BM MSC 15PH05 first 167 104 1.8 × 10¹¹ 2.0 × 10 ¹⁰ P6 (LPS- high) BM MSC 15PH05 third 177 104 7.6 × 10 ¹⁰8.4 × 10 ⁹  P8 (LPS- high) Macrophage 4/09/16 first 159 116 6.5 × 10 ¹¹3.3 × 10 ¹²

MACsplex Analysis of Exosomal Surface Markers

Comparison of exosome surface markers from exosomes from eitherunstimulated MSC or MSCs primed with high concentration LPS. The mean ofthe most intense exosome surface markers from two different human BM MSCisolates is shown in FIG. 2. Of the 37 markers analyzed by MACSPlex,both MSC and LPS-high exosomes showed the strongest surface markerprofile for eight markers; (listed from highest to lowest) CD44, CD63,CD81, CD146, CD29, CD105, MCSP and CD9. CD81, CD63 and CD9 are membersof a family oftetraspanins, known to on the surface of exosomes andcomplex with integrins for signal transduction.⁵ CD44 is a receptor forhyaluronic acid and can also interact with other ligands, such ascollagens, osteopontin, and matrix metalloproteinases (MMPs).⁶ ⁷ CD105has been found to be an auxiliary receptor for the TGF-beta receptorcomplex and is involved in modulating a response to the binding ofTGF-beta (1 and 3)⁸ CD29 is Integrin beta-1 which associates withintegrins alpha 1 and 2 to form integrin complexes that form collagenreceptors and function in a variety of processes such as tissue repair.⁹MCSP, is a cell surface proteoglycan thought be functionally importantin epidermal stem cell clustering. (Legg, J. 2003)

Comparison of Gene Expression by qPCR of Educated Macrophages vs DirectM1 Stimulation

We next examined gene expression levels that were previously found to bechanged in MEMs when co-cultivated with MSCs from different tissues.⁴⁴Gene expression in EEMs, LPS-low-EEMs and LPS-high-EEMs were compared toexpression levels in control macrophages normalized to a value of 1. Ingeneral, macrophages educated with the exosomes were moreimmunosuppressive and anti-inflammatory. There were significantincreases in expression of IDO, a known immunosuppressive modulator, inthe EEMs, along with significant increases in IL-6, IL-1B and IL-8. Verylarge increases in IDO were seen in both the LPS-low and LPS-high EEMs;increases being significant in the latter (Table 2). Increases in immunemodulating and hematopoietic system supportive IL-6 in the EEMs comparedto controls, were much higher in both of the LPS-EEMs. However,significant increases in anti-inflammatory IL-10 not seen in the EEMswere seen in the LPS-EEMs. There were also significant decreases inexpression of pro-inflammatory IL-12 in the LPS-EEMs, however there weresignificant increases in other pro-inflammatory cytokines such as TNF-αin LPS-low-EEMs and IL-8 in LPS-high-EEMs. There was also significantincrease in the VEGF-A, involved in angiogenesis in the LPS-high-EEMs.STAT1, involved in cytokine signal transduction, was higher in bothLPS-EEM populations. The M1 stimulated macrophages were morepro-inflammatory in nature, as expected, and generally produced an M1profile: anti-inflammatory IL-10 was significantly lower, along withVEGF-A, STAT1 and STAT3 and pro-inflammatory TNF-α and IL-23 weresignificantly higher.

As in Table 2, LPS-EEMs expressed a uniqueanti-inflammatory/immunosuppressive profile compared to controlmacrophages by RT-PCR. EEMs showed significant increases in mRNAexpression of IDO, IL-6, IL-1B and IL-8 expression. LPS-EEMs showedsignificant increases in expression of several anti-inflammatorycytokines (IL-6, IL-10), as well as IL-8, IDO, STAT 1 and VEGF-A. Therewas also a significant decrease in expression of the pro-inflammatoryIL-12. After M1 stimulation, macrophages produced statistical increasesin pro-inflammatory TNF-α and IL-23, and an accompanying decrease inIL-10. P values compared to control; *p</=0.05, **p</=0.005,***p</=0.0005.

TABLE 2 LPS-low LPS-high M1 Gene EEM EEM EEM stimulation IL-10 1.3 2.5*2.3** 0.17*** IDO 26.6*** 12991 12747* 62.3 IL-6 3.3* 331* 593 171 IL-121.4 0.4** 0.5** 1.1 Serpine-1 1.6 2.2 1.4 19.4* TGF-B 1.0 0.6 1.5 0.9VEGF-A 1.3 10.7 5.1*** 0.6* Stat3 1.0 1.8* 2.9 0.7* Stat1 0.8 2.8** 2.7*0.6* TNF 1.5 3.9*** 5.7 4.1** IL-23 1.3 2.6 7.0 2.9* IL-1B 2.7* 43.945.6 35.5 IL-8 9.3** 477 520.3* 542

Both MEMs and EEMs had a distinct surface marker profile by flowcytometry. The percentage of CD14+ cells with indicated surface markersand mean fluorescent intensity (MFI) of control macrophages compared toMEMs, EEMs and macrophages educated from exosomes from macrophages(macro-EEMs) by flow cytometry is shown in FIGS. 3A-3B. The % cells inthe MEMs was significantly higher than the control macrophages forCD206, PD-L1 (considered M2 markers), CD16, CD73 and the M1 marker,HLA-DR. In addition the MFIs for MEMs were higher for CD206, CD16 andCD73.²⁷. EEMs were found to express even higher levels of both % cellsand MFI for CD206 and PD-L1, and unlike MEMs which are lower for CD163but higher for PDL-2. In contrast to MEMs, CD16 and HLA-DR levels inEEMs were not significantly different compared to controls. The M1marker CD86 in either the MEMs or EEMs was not different from controls.Overall, both groups showed increased anti-inflammatory M2 surfacemarkers, but each had a distinct marker profile. The nine surface markerprofile of the Macrophage-EEMs was not significantly different thancontrols, except for PD-L1 (% cells).

As shown in Table 3, EEMs express a unique surface marker profile byflow cytometry compared to control macrophages by flow cytometry. Day 7macrophages isolated from at least three different human donors wereeither unstimulated (Control) or stimulated for 3 days by co-culturewith MSCs (MEM) or with exosomes from MSCs (EEM) or from macrophages(Macro-EEM). The ratio of CD14+ cells positive for each marker wasdesignated as percent (%) and the cell staining intensity of CD14+ cellsfor each marker was designated as median fluorescence intensity (MFI).The percentage of cells positive for CD206, PD-L1, CD16, CD73 and HLA-DRwas significantly higher in the MEMs compared to control macrophages.The MFI for CD206, CD16, and CD73 was also higher in the MEMs. In theEEMs, both the percentage of positive cells and the MFI for CD206. PD-L1and PD-L2 were also higher compared to controls. In contrast, both thepercentage of positive cells and the MFI for CD163 were lower in theEEMs compared to controls or MEMs. When comparing control macrophages tomacrophages treated with their own exosomes (Macro-EEMs) there waslittle or no difference in both the percentage of positive cells and theMFI, except for % PD-L1. P values were compared to control; *p</=0.05,**p</=0.005, ***p</=0.0005.

TABLE 3 HLA- Group CD163 CD206 PD-L1 PD-L2 CD16 CD39 CD73 CD86 DRControl 47.4% 53.8% 64.1%  63.5% 75.7% 50.3% 1.9% 75.2% 72.9% (1171)(1618) (3518) (2267) (17478) (1662) (1332) (12282) (20791) MEM 70.2% 86.2%** 80.7%* 81.7%  94.4%* 55.3% 13.3%  87.0% 91.2% (1491)  (3802)*(2436) (1916)  (32904)** (1056) (2054) (10973) (13452) EEM  23.0%* 80.3%*  89.7%**  87.2%** 80.4% 66.1% 2.1% 69.4 85.3%  (402)*  (6071)***  (12252)**  (4608)** (19925) (2701) (1542) (12033) (17201)Macro- 50.4% 66.2% 87.3%*   81% 82.5% 59.9% 1.0% 80.2 72.2% EEM (1662)(3424) (4980) (3160) (18715) (2872) (1334)  (9943) (24419)

LPS-High-EEMs Showed a High CD73 Marker and Low M1 Marker SurfaceProfile by Flow Cytometry.

Both the LPS-low EEMs and LPS-high EEMs each have a distinctive surfacephenotype when comparing both the % CD14⁺ cells with each surface markerand their MFI to each other or when compared to control macrophages orthe EEMs. Surface markers of control macrophages, EEMs, LPS-low-EEM, andLPS-high-EEMs for % CD14+ cells and MFI are shown in FIGS. 3A-3B.Compared to control macrophages, the LPS-low-EEM showed a hybrid M2-likesurface marker profile for % cells and MFI that were similar to bothEEMs and MEMs. As seen for the EEMs, LPS-low-EEMs showed significantlyelevated % cells for CD206, PD-L1 and PD-L2 compared to controls (FIG.3A). As with the MEMs, the LPS-low-EEMs had higher CD16 and CD73% cellscompared to controls. MFI (FIG. 3B) of the LPS-low-EEMs showed a higherCD206, PD-L1 and CD16 compared to controls. However, LPS-high-EEMsshowed a distinct surface profile compared to the LPS-low-EEMs. Thelevels of many M2 markers (CD206, PD-L1 and PD-L2) decreased in theLPS-high-EEMs compared to control macrophage levels. Furthermore, theLPS-high-EEMs compared to the LPS-low-EEMs expressed an even higherpercent of cells with CD73, an ecto-nucleotidase which converts AMP toadenosine and thought to be involved in immune-suppression. Importantly,lower levels of CD16, considered an inflammatory CD14+ cell markerinvolved in antibody dependent cell mediated cytotoxicity, was found inthe LPS-high-EEMs. As found for the LPS-low-EEMs, there was asignificant decrease in the M1 markers, CD86 and HLA-DR in theLPS-high-EEMs. Overall, the distinguishing marker profile of theLPS-high EEM compared to control expressed a unique surface profilewhich would be summarized as CD73-high, CD316-low, CD86-low andHLA-DR-low as shown in FIGS. 3A-3B.

As depicted in FIGS. 3A-3B, LPS-EEMs express high levels of CD73(ecto-5-nucleotidase) but low levels of M1 markers CD86 and HLA-DR byflow cytometry. Day 7 macrophages isolated from at least three differenthuman donors were either unstimulated (Control) or stimulated for 3 dayswith exosomes from MSCs (EEM) or from MSCs primed with low (LPS-low EEM)or high (LPS-high EEM) LPS. The ratio of CD14+ cells positive for eachmarker was designated as percent (%) and the cell staining intensity ofCD14+ cells for each marker was designated as median fluorescenceintensity (MFI). The LPS-low EEMs showed a higher percentage of cellspositive for CD206, PD-L1, PD-L2, CD16 and CD73 but much lower levels ofthe M1 markers, CD86 and HLA-DR compared to control macrophages. Incontrast to the LPS-low EEMs, the LPS-high EEMs marker profiles forCD206, PD-L1, PD-L2 and CD16 were low but there were significantly moreCD73 expressing cells in the LPS-high EEMs coupled with very low levelsfor CD86 and HLA-DR.

LPS-High-EEMos Showed a High PD-L1, (C73 Marker and Low M1 MarkerSurface Profile by Flow Cytometry.

The LPS-high-EEMos had a distinctive surface marker phenotype, asdetermined by flow cytometry, when compared to controls monocytes or toEEMos. As shown in FIG. 4, the flow profile (percent cells) of theLPS-high-EEMos showed high PD-L1 and CD73 expression but low CD206,CD16, PD-L2, CD163 and CD86 compared to control monocytes. When theLPS-high-EEMos were compared to the EEMos, their profile was moresimilar than controls but the levels in LPS-high-EEMos weresignificantly lower for CD16. CD73, CD86 and CD206 but higher for PDL-1.

Comparison of Gene Expression by qPCR of EEMos

As shown in FIGS. 5A-5C, gene expression studies by qPCR of theLPS-EEMos, when compared to both the control monocytes (value of 1.00)and EEMos, showed statistical increases in IL-6, IDO, FGF2, IL-10, andIL-15. IL-8 was high in the LPS-EEMos but not statistically significant.VEGF-A expression was statistically higher in the LPS-EEMos compared toonly the control monocytes. The EEMos showed statically higherexpression in IDO, but lower expression of IL-15 and IL-10 compared tocontrols. High expression of IL-6, IL-10 and IDO were common in botheducated macrophages and monocytes.

Anti-Inflammatory Immunosuppressive and Reparative Secretion Profile inthe LPS-High EEMs by Multiplex ELISA

The multiplex ELISA data indicated that macrophages educated withLPS-EVs produced from MSCs activated a large cascade of many cytokines,chemokines and growth factors that were secreted at significantly higherlevels than control macrophages. Moreover, many were significantlyhigher that the unprimed EVs used to produce the EEMs.

As shown in Table 4, LPS-EEMs secrete high levels of anti-inflammatory,growth and chemotactic factors by ELISA compared to control macrophages.Day 7 macrophages were either unstimulated (Control) or stimulated for 3days MSC-EVs to produce (EEEM) or from MSC-EVs primed with low or highLPS (LPS-low EEM) (LPS-high EEM). The EEMs secreted significant levelsof Eotaxin, G-GSF, FRACTALKINE, INFa2, GRO, IL-7, IL-8, TNF-α, and VEGFcompared to control macrophages. While not statistical due tovariability between macrophage isolates, high levels of secreted IL-6were also detected in the EEMs. When comparing EEMs to the LPS-low EEMand LPS-high EEMs, there was a significant increase in secretion of manyanti-inflammatory cytokines such as IL-4, IL-10, IL-13, including veryhigh, although non-statistical, increases in immune-modulating IL-6. Inaddition, there were significant increases inchemotactic/chemoattractant chemokines; EOTAXIN, IL-8, GRO and IP-10,growth factors such as EGF, FGF-2, and VEGF, in the cell proliferativecytokine, IL-15, soluble CD40 ligand (sCD40L) a marker of plateletactivation, hematopoietic growth factors; IL-7, platelet-derived growthfactor, two B unity type (PDGF-BB) and FMS-like tyrosine kinase type 3ligand (FLT-3L) involved in activating hematopoietic progenitors andhematopoietic cell mobilization factors, G-CSF and GM-CSF andimmunomodulatory cytokines; INFa2, IFNg, IL-17, IL-1a, IL-9 and IL-5.Both the LPS-low and -high EEM also secreted higher levels ofpro-inflammatory cytokines such as TNF-α, IFNg, IL-1b and IL-12p40 andp70. When comparing any statistical differences in secreted factorsbetween LPS-low EEMs to LPS-high EEMs, FLT-3 and IL-1, both involved incell activation and proliferation, were found to be significantly higherin the latter.

TABLE 4 Symbol (*) indicated significance versus control, (#) indicatessignificance versus EEMs, and ($) indicates significant versus ofLPS-high EEMs compared to LPS-low EEMs. The number of symbols (*, #, S)indicates the increased level of significance, one symbol p </= 0.05,two symbols p </= 0.01 three symbols p </= 0.001, 4 symbols p </= 0.0001Analyte (pg/ml) Control EEMs LPS-low-EEMs LPS-high-EEMs EGF 0.6 0.0  4.7*** ###   5.6**** #### FGF-2 17.6 21.4  34.4*** ###  35.7**** ###EOTAXIN 3.4 5.6*   8.7*** #   9.6*** ## TGF-a 2.7 3.3   6.2* #   6.4* #G-CSG 26.5 62.4*  223.9* #  286.0* ## FLT-3L 8.7 9.4  14.4* #  18.9***###$ GM-CSF 9.0 12.0  20.2** #  24.1** ## FRACTALKINE 20.1 30.2*  50.2**#  59.0*** ## INFa2 15.3 21.8*  38.5*** ##  41.1*** ### IFNg 5.8 7.6 12.1*  14.3** ## GRO 307.1 1249.6* 3317.7* 4733.3** # IL-10 38.4 45.0 500.1 1006.2** ## MCP-3 190.7 288.2  548.4  631.1* IL-12p40 8.5 9.5 18.2* #  21.4** ## MDC 4826.0 5229.0 4904.0 3517.3 IL-12p70 2.6 3.5  5.6** #   5.9**# IL-13 4.4 3.9   6.2   6.8# PDGF-BB 90.4 117.8  457.3*#  447.5* # IL-15 2.0 2.2   5.3**** ####   6.2**** ####$ sCD40L 5.6 7.0 18.7*** ###  21.5*** ### IL-17 1.1 1.6   2.7** #   2.8** # IL-1ra 820.3671.0  510.6  454.9 IL-1a 0.0 0.36  24.2*** ##  33.9*** ### IL-9 0.5 0.4  1.9   2.4* # IL-1b 1.6 2.1   5.7*   7.7** ## IL-2 1.8 2.1   3.2   3.6*IL-4 17.4 32.8  64.4*** ##  72.4*** ## IL-5 0.0 0.0   1.3* #   0.97 IL-60.0 35.6  309.9  348.2 IL-7 4.5 8.6*  23.3** #  25.5** ## IL-8 159.92251.0* 7139.0** # 8227.3*** ## IP-10 42.7 53.6 1914.3* # 2912.7** ##MCP-1 8567.7 9087.7 8983.7 8757.3 MIP-1a 18.8 36.9  145.4 1161.0* MIP-1b36.6 125.0  504.4  904.3* # RANTES 14.3 33.2  306.6  291.8 TNFa 4.014.4*  166.7* #  250.2** ## TNFb 0.0 0.0   1.4   1.7 VEGF 27.4 47.3*  81.8*** #  92.0*** #

Increased Phagocytic Activity in the LPS-High EEMS

There was a significant increase in the percentage of cells containingpHrodo Green E. coli particles in the LPS-high EEMs compared tocontrols, EEMs and macrophages directly stimulated with M1 factors (FIG.6) However, the amount of pHrodo Green E. coli particles within eachcell (MFIs) was not statistically different between all of the groups(data not shown). While macrophages (CD14⁺CD90⁻) co-cultured withMSCs(MEMs) gated with CD14 and CD90 showed a general increase inpercentage of cells with phagocytic activity, the MFI was highestoverall in this group and statistically higher than the EEMs and the M1stimulated macrophages. Overall, the only group that had a significantincrease in percentage of phagocytic cells were LPS-high EEMs.

LPS-EEMs Suppress In Vitro T-Cell Proliferation in the ImmunopotencyAssay (IPA).

The ability of cells to immuno-modulate T-cell proliferation in vitro isthought to be predictive of efficacy in vivo. Using the IPA assay, asshown in FIGS. 7A-7B, we tested the ability of macrophages, EEMs orLPS-low EEMs to modulate the growth of primary T-helper cells (CD4+) orT-cytotoxic cells (CD8+) from peripheral blood mononuclear cell (PBMCs)when co-cultured at various ratios. The LPS-low EEMs compared to theother macrophage test groups (control macrophages or EEMs) were the mostimmune-suppressive to both T-cell types. MSCs were also tested in thisassay and served as positive control inhibitor. Strongimmuno-suppression of T-cell growth by MSC has been well-documented(Bloom, D. et al. Cytotherapy, 2015, 17(2) 140-151) and MSCs were foundto be very effective at suppressing proliferation of both CD4+ and CD8+cells in the IPA assay. Almost 100% growth suppression for both CD4+ orCD8+ cells using MSCs at the 1:1 and 1:0.5 ratio while about 10-15%suppression at a 1:0.1 ratio occurred as shown in FIGS. 7A and 7B,respectively. At 1:1 and 1:0.5 there were relative degrees of a celldose-dependent suppression of PBMC proliferation when comparingmacrophage groups for suppression (i.e., control macrophages, EEMs orLPS-low-EEMs). The strongest suppression of proliferation for both CD4+and CD8+ cells occurred using the LPS-low-EEMs. While proliferationusing either macrophages or EEMs was marginally reduced about 55%proliferation for both CD4+ and CD8+ cells, there was only about 10%proliferation for both T-cell types for LPS-low-EEM ratio. IPA resultsdemonstrate the immune-suppressive properties of LPS-low EEMs on CD4+and CD8+ proliferation.

LPS-High EEMs Protect Mice from Lethal Radiation Injury in Part byRestoring Hematopoiesis.

After a lethal dosage of radiation at 4 Gy, a single intravenoustreatment with MSCs or control-macrophages did not significantly protectagainst radiation injury compared to PBS treated controls and 100% ofthese mice died within 16 days. In contrast, either EEMs or LPS-low EEMssignificantly improved mean survival from 10.6 days in the PBS controlmice to 13.2 and 18.1 days, respectively (FIG. 8A). However, LPS-highEEMs treatment led to a sustained and prolonged survival with asignificant improvement in mean and median survival of 40.7 days and47.5 days, respectively. Unlike the other treatment groups, while meanpercent weight change (FIG. 8B) and mean clinical score (FIG. 8C)worsened with time, LPS-high EEM treated mice temporarily recovered byDay 10 with clinical scores that retained somewhat normal for manyweeks. For example, the mean clinical score in LPS-high EEM treated miceout to Day 40 post-challenge remained under 2.0 compared to other groupswhich ranged from 3.0 to 5.0 during the first week. While thisprotective effect was strong, it began to diminish starting at about Day40 and the cumulative clinical score and weight progressively got worseand the remaining mice died on Day 48-52. This effect was seen after atleast three independent studies.

To determine the effects of radiation on hematopoiesis, whole blood wasassayed to determine complete blood counts (CBC) in the PBS, EEM andLPS-high EEM treated mice after lethal radiation exposure. Mean valueswere determined for three hematology populations: erythrocytes,leukocytes and thrombocytes (Table 5A, 5B, and 5C). While NSG mice areknown to be deficient in T, B and NK cells, there were detectable levelsof lymphocytes in the blood. Mice were first bled 4 days post-challengeand compared to age matched un-irradiated mice (N=10) as healthycontrols. In general, by day 4, all irradiated groups developedpancytopenia as compared to healthy controls. For erythrocytes (Table5A), the drop in cell numbers were small but significant due to thesmall variation between samples within each group. At Day 4, there weresignificant reductions in most leukocyte subsets (neutrophils,lymphocytes) in the treatment groups (Table 5B), although they remainedgenerally higher in the PBS group. Interestingly, the greatestsignificant drop in leukocyte cell subsets was detected in LPS-high EEMstreated mice, especially for neutrophils, lymphocytes and monocytes. Asignificant and approximately equal 3-fold reduction in platelets wasseen in all treatment groups (Table 5C).

Table 5, 5A, 5B, and 5C: LPS-EEM treatment significantly improved thecomplete blood count in mice after challenge with lethal radiation.(A-C) On day 0, NSG mice received 4 Gy of lethal radiation followed byan i.v. treatment 4 hours later with PBS, or with 10⁶ cells of EEMs,LPS-low or high EEMs generated as described in Methods. Blood fromfifteen non-irradiated age-matched NSG mice (normal control) wascollected to serve as the source for the normal control baseline CBCvalues. Blood was harvested from 5-10 mice from each group at 4 dayspost radiation challenge and on day 32 and days 50-53 from survivors inthe LPS-high-EEM group. The whole blood as analyzed within several hourson a Hemavet 950FS blood analyzer. The CBC values measured for the majorblood cell types from subjects in the different treatment groups areshown in Table 5. CBC values for other hematologic panel markers foreach of the three major blood groups (erythrocytes, leukocytes, andthrombocytes) are shown in Tables 5A, 5B, and 5C. Table 5A shows meanblood values of the erythrocyte panel, Table 5B shows mean blood valuesof the leukocyte panel, and Table 5C shows mean blood values of thethrombocyte panel. P values were compared to normal control mice*p</=0.05, **p</=0.005, ***p</=0.0005.

TABLE 5 CBC values for major blood cell types (mean values) Day Plateletpost RBC WBC Neutrophils Lymphocytes Monocytes Platelets vol Groupradiation (M/ul) (K/ul) (K/ul) (K/ul) (K/ul) (K/ul) (fL) Control N/A 4.61.37 1.06 0.21 0.065 608  4.5 PBS 4 4.3 0.49* 0.14** 0.24 0.03 191** 4.3EEM 4 3.7 0.19** 0.03** 0.08 0.013 219** 4.2* LPS- 4 3.7* 0.21***0.02*** 0.05* 0.01**  187*** 4.5 high EEM LPS- 32 4.1 1.68 1.21 0.360.05 379*  5.0*** high EEM LPS- 50-53 5.8 1.48 0.92 0.40 0.11 316**5.0*** high EEM

TABLE 5A Erythrocyte panel (mean values) Day post RBC Hb HCT MCV MCHMCHC RDW Group radiation (M/ul) (g/dL) (%) (fL) (pg) (g/dL) (%) NormalN/A 4.6 6.2 22.8 48   13.6 28.2 16.8 control PBS 4 4.3 5.8 20.1 46.3*13.3 28.7 15.9** EEM 4 3.7 4.8* 17.2 46.6* 12.9* 27.8 15.6*** LPS- 43.7* 5.0* 17.6*  46.7** 13.3 28.5 15.9*** high- EEM LPS- 32 4.1 5.4 21.6  52.2*** 13.5 26.1* 21.1*** high EEM

TABLE 5B Leukocyte panel (mean values) Day post WBC NeutrophilsLymphocytes Monocytes Eosinophils Basophils Group radiation (K/ul)(K/ul) (K/ul) (K/ul) (K/ul) (K/ul) Normal N/A 1.37 1.06 0.21 0.065 0.0310.014 control PBS 4 0.49* 0.14** 0.24 0.03 0.04 0.03 EEM 4 0.19** 0.03**0.08 0.013 0.01 0.003 LPS- 4 0.21*** 0.02*** 0.05* 0.01** 0.006 0.003high EEM LPS- 32 1.68 1.21 0.36 0.05 0.04 0.011 high EEM

TABLE 5C Thrombocyte panel (mean values) Group Day post radiationPlatelets (K/ul) Platelet volume (fL) Normal control N/A 608 4.5 PBS 4191** 4.3 EEM 4 219** 4.2* LPS-high EEM 4 187*** 4.5 LPS-high EEM 32379* 5.0***

The levels of most blood cell types of all three hematology panels wererestored to normal levels by Day 32 in the surviving LPS-high EEM mice.Accordingly, most of these mice displayed near normal weight andclinical scores (FIGS. 8A-88). For erythrocytes, most of the values wentto normal levels with MCV (mean corpuscular volume) and RDW (red celldistribution width) significantly higher than normal controls, likelyfrom increased reticulocytosis. Both of these parameters indicatestimulation of red cell production by the LPS-high EEMs. Furthermore,the mean values for leukocyte subsets were all restored to normal levelsby Day 32. While platelets did not reach normal levels, amountssignificantly improved, (Table 5C), with also significantly higherplatelet volume indicating increased production of immature plateletsfrom the BM hematopoietic progenitor cells. Significantly the CBC panelperformed on the relapsed moribund LPS-high EEM treated mice (Day 50-53)was still very similar to the CBCs of the recovered healthy Day 32LPS-high EEM treated mice.

In another lethal radiation study, mice from different treatment groupswere euthanized at key time points post challenge for gross necropsy andwere examined by histology to determine the status of theirhematopoietic organs (bone marrow and spleen) and possible cause ofdeath. Healthy, un-irradiated NSG mice served as normal control tissuewhile several sets of mice were irradiated at 4 Gy and treated witheither PBS (control) or LPS-High EEMs as described. Irradiated PBScontrol mice showing overt signs of ARS (Day 9) were euthanized and adetailed gross necropsy was conducted including organ weight/morphologyand histology and compared to healthy control tissue. Two sets ofLPS-High EEM treated mice were also compared—the healthy Day 30 mice andthe Day 50-53 relapsed mice. Spleen weight can be used as reliablemarker for presence or lack of extra-medullary hematopoiesis. As shownin Table 6 below, there was a significant drop in both spleen weight andspleen % body weight (BW) in the irradiated untreated mice compared tothe spleens of normal mice. In contrast, mean spleen weights weresimilar to normal in the LPS-High EEM treated mice at Day 30post-challenge. However, the LPS-High EEM treated mice at Day 50-53showed clinical signs of relapse and weight loss including significantlylower mean spleen weights and spleen % BW as compared to those valuesobtained from healthy control mice.

TABLE 6 Treatment group Spleen weight (mg) Spleen % BW Normal 28.1 0.114GY untreated (Day 8) 9.3* 0.05* 4GY LPS-High EEM (Day 34.4 0.15 30) 4GYLPS-High EEM (Day 12.3* 0.07* 50-53) *P = > 0.05

The spleen sizes were also examined across the treatment groups. Day 8irradiated, untreated mice compared to the normal mice indicated therewas clear histopathology present with absence of hematopoietic cells(both progenitor and mature cells types) in both the spleen and bonemarrow. This was clearly reflected in the profound reduction seen in theCBCs (Table 5). In contrast, spleen size of the LPS-high Day 30 miceshowed healthy and prominent active hematopoietic tissue in the bonemarrow and spleen, and again also reflected in a re-established CBC(Table 5). Interestingly, the histopathology of the blood, spleen andbone marrow of Day 50-53 moribund LPS-high EEM mice still remainedfairly unremarkable, and all showed hematopoietic tissue in the bonemarrow and marked extra-medullary hematopoiesis in the spleen. Eventhough the mean spleen size was reduced in these mice, signs of both anormal CBC and spleen and BM histology indicate that the moribundLPS-high EEM mice after Day 50 most likely did not die from severeanemia (low RBC counts) or leukopenia (low white cell count) but fromanother undetermined reason. Therefore, it appears that a singleinjection of LPS-high EEMs can restore a functioning hematopoiesis inthe bone marrow and the spleen in the mice long term after challengewith lethal radiation and that death in these mice may not be due to theloss of radioprotection in these tissues.

LPS-High-EEMs Protect Mice from Lethal Radiation Injury by RestoringHematopoietic Tissue.

To identify which organs and tissues may be protected by LPS-high EEMtreatment, we compared histology of BM from femur (FIG. 9A) and spleens(FIG. 9B) of normal non-irradiated mice to irradiated mice with orwithout treatment at different times post-challenge. By gross necropsy,the spleens were most affected by 4 Gy radiation exposure, while overtchanges to the heart, liver and kidneys were less obvious (data notshown). Compared to the histologic sections of BMs and spleens fromhealthy mice (FIGS. 9A and 9B), moribund PBS treated mice at day 9post-irradiation showed a marked absence of hematopoietic cellularity inthe BM and total lack of extra-medullary hematopoiesis in the spleenwith clear hemorrhage (FIGS. 9A and 9B, respectively). In contrast,there were markedly more hematopoietic cells present in the LPS-EEMtreated mice at day 9 post-irradiation (FIGS. 9A and 9B). At this time,cellularity present in the BM cavity graded from 1 to 5 (indicating mostto least) was only 4 to 5 in the untreated mice post-radiationchallenge, but ranged from 0.5 to 3.0 in the LPS-high EEM mice.Improvement continued in these mice at day 30 with strong to moderatehematopoietic activity in the BM of the femur but also in the pelvis andsternum with an intense hematopoietic component present in spleen.Interestingly even during clinical symptom relapse at day 53,hematopoietic tissue in the BM and spleen was still distinctly presentin the LPS-high EEM treated mice, similar to what we observed in CBCs.

LPS-High EEMos Protect Mice from Lethal Radiation Injury

After a lethal dosage of radiation at 4 Gy, a single intravenoustreatment with control monocytes or EV (exosome) educated monocytes(EEMos) did not significantly protect against radiation injury comparedto PBS treated controls and 100% of these mice died within 12 days. Incontrast, LPS-high EEMos treatment led to a sustained and prolongedsurvival with a significant improvement, all mice survived for 45 days.Unlike the other treatment groups, while mean percent weight change(FIG. 8B) and mean clinical score (FIG. 8C) worsened with time, LPS-highEEM treated mice temporarily recovered after Day 10 and both weights andclinical scores remained normal for many weeks until about Day 40. Asalso seen in the treatment studies with the LPS-high EEMs, after asingle treatment the effects of the LPS-high EEMos began to diminishstarting at about Day 40 and the cumulative clinical score and weightprogressively got worse and the mice died on Day 45-49.

To determine the effects of radiation on hematopoiesis, whole blood wasassayed to determine complete blood counts (CBC) in the PBS, EEMos andLPS-high EEMos treated mice after lethal radiation exposure. Mean valueswere determined for three hematology populations: erythrocytes (RBCs),leukocytes (WBC, neutrophils, lymphocytes and monocytes) andthrombocytes (platelets and platelet volume) (Table 7). Blood fromnon-irradiated age-matched NSG mice (normal control) was collected toserve as the source for the normal control baseline CBC values.LPS-EEMos treatment significantly improved the complete blood count inmice after challenge with lethal radiation. Blood was harvested from themice from each group at 5 days post radiation challenge and on day 30from the surviving members in the LPS-high EEM group. The whole bloodwas analyzed within several hours on a Hemavet 950 FS blood analyzer. Asseen in the CBC results in the same animal model using LPS-high EEMs, atDay 5 the CBC dropped significantly for most of the cells types in eachhematology population. However, CBCs significantly improved to normallevels, except platelets (which did improve to near normal levels) atDay 30 and even Day 48, when the mice relapsed and were moribund.

TABLE 7 CBC values for major blood cell types (mean values) Platelet Daypost RBC WBC Neutrophils Lymphocytes Monocytes Platelets volume Groupradiation (M/μl) (K/μl) (K/μl) (K/μl) (K/μl) (K/μl) (fL) Normal N/A 8.64.31 2.13 1.63 1.63 954.0 4.8 Control PBS 5 8.6 0.45*** 0.11*** 0.29*0.03*** 298.0** 4.8 EEMos 5 8.3 0.36*** 0.06*** 0.23* 0.04*** 340.0***4.4 LPS- 5 8.7 0.53** 0.13*** 0.41 0.41** 316.0*** 4.9 high- EEMos LPS-30 7.46* 2.73 1.19 1.23 0.125 498.45*** 5.23* high- EEMos LPS- 48 9.814.79 2.42 1.97 0.20 691.67* 4.87 high- EEMos *p </= 0.05, **p </= 0.005,***p </= 0.0005

Exosome Dose Response Studies Indicate that LPS-High Exosomes can AlsoProtect Mice from Lethal Radiation Injury

When the same concentration of LPS-high exosomes used to educate cells(either 10⁶ macrophages or 107 monocytes) and successfully treat micewas used to directly treat mice, that dose was unable to significantlyprotect against death from radiation injury compared to PBS treatedcontrols (FIG. 11.) Specifically after a lethal dosage of radiation at 4Gy, mice treated directly with a single intravenous treatment of 2.5×10⁹exosomes from either unstimulated MSCs or exosomes from MSCs stimulatedwith LPS (high) exosomes were not effective in protecting fromlethality. In contrast, as shown previously at this same dose, educationof either monocytes or macrophages with LPS-high exosomes wassuccessfully able to generate cells that were protective in theradiation model. This indicates that this exosome dose is enough toeffectively educate cells protective for lethal radiation injury inmice, but not enough to be effective when used directly.

Increasing the number of exosomes for direct use in the radiation modelwas then tested. Exosomes from either unstimulated MSC exosomes orLPS-high exosomes at a two-fold increase to 5.0×10⁹, producedsignificant improvement in mean survival for both the MSC-exosome andthe LPS-high exosome treated mice compared to the PBS control mice (FIG.12A). When LPS-high exosomes at this higher dose were also used toeducate monocytes to generate LPS-high EEMos, significant improvement insurvival compared to control mice was also seen as expected (FIG. 12A).This higher exosome dose to generate LPS-high EEMos did not appear toimprove either clinical outcome or prolong survival significantlycompared with education using a lower dose (see FIG. 10A). As witheither LPS-high EEMs or LPS-high EEMos, mice treated with high doseexosomes also showed significant improvement in both the mean clinicalscore (FIG. 12B) and mean % weight change. (FIG. 12C). As seenpreviously using LPS-high EEMs or LPS-EEMos both the clinical score and% weight changes worsened with time starting at day 40 in mice treateddirectly with higher dose MSC or LPS-high exosomes and all of the micedied by about day 50. However is noteworthy that the clinical scores inthe mice treated with LPS-high EEMos were significantly better comparedto directly treating with either set of exosomes (FIG. 12B).

High Dose Exosomes Treatment can Help Maintain Normal Complete BloodCounts in Mice after Lethal Radiation Injury

To determine the effects of radiation and high dose exosome treatment onhematopoiesis, whole blood collected from the mice (FIG. 12) was assayedto determine complete blood counts (CBC) in the PBS, MSC exosome,LPS-high exosome, and the LPS-high EEMo treated mice after lethalradiation exposure as before. Mean values were determined for three keyhematology populations: erythrocytes (RBCs), leukocytes (WBC, and types:neutrophils, lymphocyte and monocytes) and thrombocytes (platelets,platelet volume) (Table 8). Mice were first bled 4 days post-challengeand compared to non-irradiated mice as normal healthy controls. Ingeneral, by day 4, all irradiated groups developed pancytopenia ascompared to healthy controls. As seen in previous studies at this timepoint (day 4-5), there were significant reductions in cell numberswithin all groups; specifically WBC, neutrophils and platelets.Interestingly, there was also a significant increase in platelet volumeat this early time point in both the LPS-high exosome and LPS-high EEMosnot typically seen in earlier studies using LPS-high EEMos educated atthe lower doses of exosomes. An increase in platelet volume typicallyindicates active proliferation of platelets. By day 30, the WBC andplatelets recovered in the MSC exosome treated mice to normal levels butneutrophils remained lower and lymphocytes were statistically higher,both outcomes typically not seen in earlier studies with LPS-high EEMsand EEMos. In contrast the CBC levels in both LPS-high exosomes and theLPS-high EEMos returned to normal by day 30, except platelet volumeremained higher in the LPS-high EEMos treated group. CBC determinationsof blood samples from moribund mice at day 44 treated with LPS-highexosomes or LPS-high EEMos indicate that the hematology populations werestill near normal compared with controls; although the WBC, neutrophilsand platelet volume were significantly higher, but still within thenormal healthy mouse range. The CBC of the surviving MSC-exosome treatedmice at this time was similar to both LPS-high exosomes and LPS-highEEMos, except the platelet levels dropped below normal controls. Theresults indicate that the CBCs in mice treated with MSC exosomes,LPS-high exosomes or LPS-high EEMos recover after lethal radiation andthat even during clinical relapse, the CBC levels still remain largelynormal indicating a still functioning hematopoietic system.

Table 8 demonstrates that using higher dosages of MSC-exosomes, LPS-highexosomes and LPS-high EEMos significantly improved the complete bloodcount in mice after challenge with lethal radiation. On day 0. NSG micereceived 4 Gy of lethal radiation followed by an i.v. treatment 4 hourslater with PBS, or 5×10⁹ MSC-exosomes or LPS-high exosomes or 107monocytes treated with 5×10⁹ LPS-high exosomes producing LPS-high EEMos.The educated monocytes were generated as described in Methods. Bloodfrom non-irradiated age-matched NSG mice (normal control) was collectedto serve as the source for the normal control baseline CBC values. Bloodwas harvested from surviving members from each group at day 4, day 30and day 44 post radiation challenge. The whole blood as analyzed withinseveral hours on a Hemavet 950FS blood analyzer. (A) Mean blood valuesof the erythrocyte, leukocyte panel, and thrombocyte panel is shown. Pvalues were compared to normal control mice *p</=0.05, **p</=0.005,***p</=0.0005.

TABLE 8 CBC values of major blood cell types (mean values) Platelet Daypost RBC WBC Neutrophils Lymphocytes Monocytes Platelets volume Groupradiation (M/ul) (K/ul) (K/ul) (K/ul) (K/ul) (K/ul) (fL) Normal n/a 9.022.45 1.63   0.57  0.13  794.20 5.03   Control PBS 5 8.64    0.507 *** 0.140 *** 0.293 0.037    205.33 ** 5.0    MSC 5 8.39    0.28 ***0.037*** 0.137 0.023    195.67 ** 4.97   Exosomes LPS-high 5 10.36   0.167 ***  0.020 *** 0.053 0.017    150.3 *** 6.3 *** ExosomesLPS-high 5 9.08    0.35 *** 0.12***  0.193 0.017   204.0 ** 6.1 ** EEMos MSC 30 7.46 2.38 0.835*    1.265 * 0.2 880.0 5.75 ** ExosomesLPS-high 30 8.00 1.28 1.01   0.23  0.03 628.5 5.5    Exosomes LPS-high30 8.40 2.65 0.99   1.14  0.04  506.25 6.15 ** EEMos MSC 44 6.86  4.09 * 2.93 *  0.75  0.23   430.5 * 5.9 **  Exosomes LPS-high 44 9.94  4.42 ** 4.09 *** 0.23  0.06 567   5.85 *  Exosomes LPS-high 44 8.56  3.44 * 2.82 **  0.45  0.11 790   5.63 ** EEMos

Discussion

Presently protection from radiation injury involves mainly supportivecare and treatment with growth factors until an allogeneic BMT can bearranged. Development of cell-based therapies for radiation injury areappealing because of the potential to infuse them soon after radiationinjury, produce multiple cytokines that can protect multiple organs fromtissue injury and potentially restore hematopoiesis. Macrophages arelong-lived phagocytic cells that differentiate from circulatingmonocytes and migrate into tissues to replace older cells or in responseto signals from tissue injury. Once at the site of injured tissue,macrophages are an essential component in the host defense by clearingthe site of pathogens, regulating inflammation, and promoting tissuerepair.⁴⁷ In this environment, macrophages are very plastic and respondto products released from damaged tissue or pathogens (e.g. Toll-likereceptors, LPS) to become microbicidal or M1-like. Local MSCs also sensethese inflammatory products and respond to restore homeostasis bycommunicating with effector cells such as macrophages or T-cells throughparacrine factor and exosomes⁴⁸⁻⁵⁰ or direct contact²⁷ to promote analternative M2 or T-regulatory phenotypes to orchestrate tissue repair.Direct exposure of macrophages to microbial factors such as LPS canpolarize them to M1-like macrophages (M1 stimulation) while exposure ofexosomes from LPS-primed MSCs can lead to the induction of reparative ormore M2 like macrophages. The goal of this study was to determine ifMSC-derived exosomes could be used to educate macrophages and monocytesinto a radio-protective phenotype. In vivo bacterial sepsis modelsindicate MSCs help increase the percentage of detectable M2macrophages⁵¹ and LPS-primed MSCs are better at promoting tissuehealing.⁵² Furthermore, a recent study showed that exosomes fromLPS-primed MSCs show improved wound healing in diabetic rat model.³⁷Thus an interesting paradox exists where the same inflammatory mediator,such as LPS, can induce different downstream responses depending onwhich cell type it encounters.

Interestingly while LPS-high-EEMs showed the best radioprotection in ourlethal radiation injury model, these macrophages did not fit the typicalM2-like reparative phenotype. Based on cell surface marker and secretionprofiles, M2 macrophages can be further classified into at least 4subsets (M2a,b,c,d).⁵³ Which current subset, if any, might be mosteffective at treating radiation injury is unclear. Expression of CD163,CD206, CD274 (PD-L1) and CD273 (PD-L2) decreased compared to the EEMs,LPS-low EEMs and MEMs and essentially reversed to normal macrophagelevels. However, like MEMs also effective in the radiation model, strongCD73 ecto-nucleotidase expression likewise occurred in the LPS-EEMs.²⁷Increased adenosine production, along with huge increases in IDOexpression (the enzyme involved in T-cell suppression by tryptophandegradation)⁵⁴ along with low expression of CD16 and M1 markers CD86 andHLA-DR indicate that T-cell suppression may be very important mechanismof action in the LPS-high EEMs. Besides this immunosuppression andimmunomodulation, the LPS-high EEMs cells also have stronganti-inflammatory characteristics with increased phagocytosis aiding inthe rapid clearance of cell debris.

The LPS-EEMs secreted significantly higher levels of a variety ofanti-inflammatory cytokines, chemotactic factors, and growth factorscompared to either control macrophages or EEMs. IL-6, a hallmarkbiomarker for MEMs, has been described to induce alternatively activatedmacrophages⁵⁵, promote mucosal healing from colitis⁵⁶, and cartilageself-repair by MSCs⁵⁷ and decreased ICAM-1 secretion, both known toreduce radiation-induced inflammation. ^(24,58) LPS-high-EEMs alsosecreted significant levels of other cytokines with anti-inflammatoryactivities such as, IL-4, L-10 and IL-13. However, there were alsosignificant increases in levels with pro-inflammatory tendenciesincluding TNF-α, IL-1b, IFN-g, IL-12p40-p70, IL-15 and IL-17. However,except for TNF-α, in general the fold increase in many of them were notat the same level as found for many of the anti-inflammatory cytokines.The large increase in chemotactic factors such as MCP-3, IL-8, and IP-10was also seen and may attract other monocytes, macrophages andneutrophils to site to treat radiation injury. The key to effectivenessof the LPS-EEMs might be due to the increases in growth factors such asEGF, FGF and G-CSF and GM-CSF. Indeed, the latter two growth factors areused clinically to treat radiation associated illness. Since theLPS-high-EEMs were more effective than the LPS-low-EEMs, two factors,FLT-3L, a stem cell growth factor which stimulates the growth of bloodprogenitors and IL-15 a regulator of proliferation of T-cells foundsignificantly elevated in the LPS-high-EEMs may indicate theirimportance for an improved therapeutic outcome seen in the radiationmouse model. Based on the secretion profile (IL-10, IL-12, TNF-alpha,IP-10), LPS-high-EEMs most resemble M2-d subtype of macrophages, howeverthere are also many unique differences such as high IL-6 and CD73 whichindicate they belong to novel and distinct subset of activatedmacrophages. LPS-high-EEMos are characterized by high expression ofPD-L1 and CD73, IL-15, and IL-6 and low expression of CD206, CD163,CD86, and CD16 compared to control monocytes.

After a lethal dose of radiation, all the animals presented with acuteradiation syndrome (ARS) indicated by significant weight loss, changesin body posture and fur texture.⁴³ Since rapidly proliferatingprogenitor stem cells in the bone marrow are most radio-sensitive, asexpected we found that all the mice had severe pancytopenia¹ and mostcell values at Day 4 in all three panels (erythrocyte, leukocyte andthrombocyte panels) were significantly lower (except lymphocyte countsin the PBS group). Acute effects of radiation injury on the BM wasmarkedly illustrated by Day 4 in all groups, but LPS-high-EEMs andLPS-high-EEMos were able to reverse most of these abnormalities by Day32 and Day 30, respectively. In addition, while the median survival inthe control groups occurred at Day 9, the majority (71%) of LPS-high-EEMtreated mice were alive at Day 32. Likewise, the majority ofLPS-high-EEMo treated mice were alive through day 40. Correspondingly,20% of the mice in the LPS-low-EEM groups were also alive at this point.At this time, cell counts of the LPS-high-EEMs of all three panelsrecovered to normal or near normal levels; especially pronounced was thesignificant restoration of the white blood count and its component cellstypes. We have previously shown that by Day 12, >50% of normal humanmononuclear cells injected i.v. were detected within the BM and spleenin irradiated mice.₂₇ Therefore, based on the overall results presentedhere, reversal of ARS in the mice by LPS-high EEM may be due in part totheir effectiveness in suppressing inflammation and restoringhematopoiesis in the BM and/or spleen. Indeed, studies have indicatedthat in response to stress, macrophages can play a role in supporting BMerythropoiesis.^(59,60)

Since survival of irradiated mice treated with a single dose of eitherLPS-low-EEMs, LPS-high-EEMs, or LPS-high-EEMos was not permanent, weoriginally thought this was due to loss of the protective effect by theLPS-EEMs or LPS-EEMos on hematopoiesis. However, subsequent in vivostudies comparing healthy Day 30 mice to moribund Day 50-53 miceindicated that both normal CBCs and functional hematopoiesis in the BMand spleen were still present in Day 50-53 mice. Interestingly,surviving LPS-EEM treated mice from three independent radiation studiesall died within a short period of time (Day 48-53). Upon necropsy andhistology, no overlying cause of death was found in these mice. Howeverthe short-time frame of death post-radiation indicate that it may not bedue to something accidental such as infection in the majority of micebut something specific such as a finite engraftment time of the humanLPS-EEM in the mice. Therefore, an additional treatment of LPS-high-EEMsat strategic time points (Day 30) when the clinical scores begin toworsen, or repeated injections before those clinical symptoms return,may result in long-term survival.

Future studies of EEMs should focus on understanding the molecularmechanisms driving LPS-high EEMs in protecting mice from lethalradiation. RNA-seq analysis of exosomes could help identify factorsimportant in the generation of radio-protective macrophages. Microarrayanalysis of microRNA (miRNA) from LPS-exosomes has showed a differentialexpression profile with elevated expression of certain miRNAs, such asthe transcription factor, let-7b; proposed to be a driver in macrophagepolarization.^(37,61) Using an unbiased RNA-seq for MEMs, expression ofgenes involved positively correlated with several pathways that could bebeneficial for anti-inflammatory effects or tissue repair (e.g. collagenformation or tissue development genes).²⁷ Testing whether survivalpost-radiation challenge can improve with increases in LPS-EEM dose andor with repeated cell treatments will be critical, as well as performinga time course to determine how long after radiation injury EEM infusioncan be administered to still contribute to survival. Overall, we hopethis work can provide the foundation to develop a more effectivetreatment of radiation injury using therapeutic macrophages andmonocytes.

REFERENCES

-   1. Williams J P, Brown S L, Georges G E, et at. Animal models for    medical countermeasures to radiation exposure. Radiat Res. April    2010; 173(4):557-578.-   2. Fliedner T M, Chao N J, Bader J L, et al. Stem cells, multiorgan    failure in radiation emergency medical preparedness: a U.S./European    Consultation Workshop. Stem Cells. May 2009; 27(5):1205-1211.-   3. Singh V K, Seed T M. A review of radiation countermeasures    focusing on injury-specific medicinals and regulatory approval    status: part I. Radiation sub-syndromes, animal models and    FDA-approved countermeasures. Int J Radial Biol. September 2017;    93(9):851-869.-   4. Singh V K, Garcia M, Seed T M. A review of radiation    countermeasures focusing on injury-specific medicinals and    regulatory approval status: part II. Countermeasures for limited    indications, internalized radionuclides, emesis, late effects, and    agents demonstrating efficacy in large animals with or without FDA    IND status. Int J Radiat Biol. September 2017; 93(9):870-884.-   5. Singh V K, Newman V L, Seed T M. Colony-stimulating factors for    the treatment of the hematopoietic component of the acute radiation    syndrome (H-ARS): a review. Cytokine. January 2015; 71(1):22-37.-   6. Koc O N, Gerson S L, Cooper B W, et al. Rapid hematopoietic    recovery after coinfusion of autologous-blood stem cells and    culture-expanded marrow mesenchymal stem cells in advanced breast    cancer patients receiving high-dose chemotherapy. J Clin Oncol.    January 2000; 18(2):307-316.-   7. Pittenger M F, Mackay A M, Beck S C, et al. Multilineage    potential of adult human mesenchymal stem cells. Science. Apr. 2    1999; 284(5411):143-147.-   8. Bernardo M E, Fibbe W E. Mesenchymal stromal cells: sensors and    switchers of inflammation. Cell Stem Cell. Oct. 3 2013;    13(4):392-402.-   9. Eaton E B, Jr., Varney T R. Mesenchymal stem cell therapy for    acute radiation syndrome: innovative medical approaches in military    medicine. Mil Med Res. 2015; 2:2.-   10. Hu K X, Sun Q Y, Guo M, Ai H S. The radiation protection and    therapy effects of mesenchymal stem cells in mice with acute    radiation injury. Br J Radiol. January 2010; 83(985):52-58.-   11. Lange C, Brunswig-Spickenheier B, Cappallo-Obermann H, et al.    Radiation rescue: mesenchymal stromal cells protect from lethal    irradiation. PLoS One. Jan. 5 2011; 6(1):e14486.-   12. Shim S, Lee S B, Lee J G, et al. Mitigating effects of hUCB-MSCs    on the hematopoietic syndrome resulting from total body irradiation.    Exp Hematol. April 2013; 41(4):346-353 e342.-   13. Hu J, Yang Z, Wang J, et al. Infusion of Trx-1-overexpressing    hucMSC prolongs the survival of acutely irradiated NOD/SCID mice by    decreasing excessive inflammatory injury. PLoS One.    2013:8(11):e78227.-   14. Wang S, Qu X, Zhao R C. Clinical applications of mesenchymal    stem cells. J Hematol Oncol. 2012; 5:19.-   15. Galipeau J. The mesenchymal stromal cells dilemma—does a    negative phase 111 trial of random donor mesenchymal stromal cells    in steroid-resistant graft-versus-host disease represent a death    knell or a bump in the road? Cytotherapy. January 2013; 15(1):2-8.-   16. Chiossone L, Conte R, Spaggiari G M, et al. Mesenchymal Stromal    Cells Induce Peculiar Alternatively Activated Macrophages Capable of    Dampening Both Innate and Adaptive Immune Responses. Stem Cells.    July 2016; 34(7):1909-1921.-   17. Nemeth K, Leelahavanichkul A, Yuen P S, et al. Bone marrow    stromal cells attenuate sepsis via prostaglandin E(2)-dependent    reprogramming of host macrophages to increase their interleukin-10    production. Nat Med. January 2009; 15(1):42-49.-   18. Cho D I, Kim M R, Jeong H Y, et al. Mesenchymal stem cells    reciprocally regulate the M1/M2 balance in mouse bone marrow-derived    macrophages. Exp Mol Med. Jan. 10 2014; 46:e70.-   19. Melief S M, Schrama E, Brugman M H, et al. Multipotent stromal    cells induce human regulatory T cells through a novel pathway    involving skewing of monocytes toward anti-inflammatory macrophages.    Stem Cells. September 2013; 31(9):1980-1991.-   20. Kim J, Hematti P. Mesenchymal stem cell-educated macrophages: a    novel type of alternatively activated macrophages. Exp Hematol.    December 2009; 37(12):1445-1453.-   21. Sica A, Mantovani A. Macrophage plasticity and polarization: in    vivo veritas. J Clin Invest. March 2012; 122(3):787-795.-   22. Keil F, Elahi F, Greinix H T, et al. Ex vivo expansion of    long-term culture initiating marrow cells by IL-10, SCF, and IL-3.    Transfusion. May 2002; 42(5):581-587.-   23. Duchez P, Rodriguez L, Chevaleyre J, et al. Interleukin-6    enhances the activity of in vivo long-term reconstituting    hematopoietic stem cells in “hypoxic-like” expansion cultures ex    vivo. Transfusion. November 2015; 55(11):2684-2691.-   24. Koukourakis M I. Radiation damage and radioprotectants: new    concepts in the era of molecular medicine. Br J Radiol. April 2012;    85(1012):313-330.-   25. Roberts C A, Dickinson A K, Taams L S. The Interplay Between    Monocytes/Macrophages and CD4(+) T Cell Subsets in Rheumatoid    Arthritis. Front Immunol. 2015:6:571,-   26. Aggarwal S, Pittenger M F. Human mesenchymal stem cells modulate    allogeneic immune cell responses. Blood. Feb. 15 2005;    105(4):1815-1822.-   27. Bouchlaka M N, Moffitt A B, Kim J, et al. Human Mesenchymal Stem    Cell-Educated Macrophages Are a Distinct High IL-6-Producing Subset    that Confer Protection in Graft-versus-Host-Disease and Radiation    Injury Models. Biol Blood Marrow Transplant. June 2017;    23(6):897-905.-   28. Caplan A I, Correa D. The MSC: an injury drugstore. Cell Stem    Cell. Jul. 8 2011; 9(1):11-15.-   29. Pittenger M. Sleuthing the source of regeneration by MSCs. Cell    Stem Cell. Jul. 2 2009; 5(1):8-10.-   30. Phinney D G, Pittenger M F. Concise Review: MSC-Derived Exosomes    for Cell-Free Therapy., Stem Cells. April 2017; 35(4):851-858.-   31. Katsuda T, Kosaka N, Takeshita F, Ochiya T. The therapeutic    potential of mesenchymal stem cell-derived extracellular vesicles.    Proteomics. May 2013; 13(10-11):1637-1653.-   32. Lai R C, Yeo R W, Lim S K. Mesenchymal stem cell exosomes. Semin    Cell Dev Biol. April 2015; 40:82-88.-   33. Yu B, Zhang X, Li X. Exosomes derived from mesenchymal stem    cells. Int J Mol Sci. 2014; 15(3):4142-4157.-   34. S ELA, Mager I, Breakefield X O, Wood M J. Extracellular    vesicles: biology and emerging therapeutic opportunities. Nat Rev    Drug Discov. May 2013:12(5):347-357.-   35. Tasso R, Ilengo C, Quarto R, Cancedda R, Caspi R R, Pennesi G.    Mesenchymal stein cells induce functionally active T-regulatory    lymphocytes in a paracrine fashion and ameliorate experimental    autoimmune uveitis. Invest Ophthalmol Vis Sci. February 2012;    53(2):786-793.-   36. English K, Ryan J M, Tobin L, Murphy M J, Barry F P, Mahon B P.    Cell contact, prostaglandin E(2) and transforming growth factor beta    1 play non-redundant roles in human mesenchymal stem cell induction    of CD4+CD25(High) forkhead box P3+ regulatory T cells. Clin Exp    Immunol. April 2009; 156(1):149-160.-   37. Ti D, Hao H, Tong C, et al. LPS-preconditioned mesenchymal    stromal cells modify macrophage polarization for resolution of    chronic inflammation via exosome-shuttled let-7b. J Transl Med.    2015; 13(1):308.-   38. Wen S, Dooner M, Cheng Y, et al. Mesenchymal stromal    cell-derived extracellular vesicles rescue radiation damage to    murine marrow hematopoietic cells. Leukemia. November 2016;    30(11):2221-2231.-   39. Dominici M, Le Blanc K, Mueller I, et al. Minimal criteria for    defining multipotent mesenchymal stromal cells. The International    Society for Cellular Therapy position statement. Cytotherapy. 2006;    8(4):315-317.-   40. Bloom D D, Centanni J M, Bhatia N, et al. A reproducible    immunopotency assay to measure mesenchymal stromal cell-mediated    T-cell suppression. Cytotherapy. February 2015; 17(2):140-151.-   41. Thery C, Amigorena S, Raposo G, Clayton A. Isolation and    characterization of exosomes from cell culture supernatants and    biological fluids. Curr Protoc Cell Biol. April 2006; Chapter 3:Unit    3 22.-   42. Sindrilaru A, Peters T, Wieschalka S, et al. An unrestrained    proinflammatory M1 macrophage population induced by iron impairs    wound healing in humans and mice. Jin Invest. March 2011;    121(3):985-997.-   43. Cooke K R, Kobzik L, Martin T R, et al. An experimental model of    idiopathic pneumonia syndrome after bone marrow transplantation: 1.    The roles of minor H antigens and endotoxin. Blood. Oct. 15 1996;    88(8):3230-3239.-   44. Kim J, Escalante L E, Dollar B A, Hanson S E, Hematti P.    Comparison of breast and abdominal adipose tissue mesenchymal    stromal/stem cells in support of proliferation of breast cancer    cells. Cancer Invest. October 2013; 31(8):550-554.-   45. Linden J. Molecular approach to adenosine receptors:    receptor-mediated mechanisms of tissue protection. Annu Rev    Pharmacol Taxicol. 2001; 41:775-787.-   46. Yang L, Froio R M, Sciuto T E, Dvorak A M, Alon R, Luscinskas    F W. ICAM-1 regulates neutrophil adhesion and transcellular    migration of TNF-alpha-activated vascular endothelium under flow.    Blood. Jul. 15 2005; 106(2):584-592.-   47. Gordon S, Martinez F O. Alternative activation of macrophages:    mechanism and functions. Immunity. May 28 2010; 32(5):593-604.-   48. Blazquez R, Sanchez-Margallo F M, de la Rosa O, et al.    Immunomodulatory Potential of Human Adipose Mesenchymal Stem Cells    Derived Exosomes on in vitro Stimulated T Cells. Front Immunol.    2014; 5:556.-   49. Zhu Y G, Feng X M, Abbott J, et al. Human mesenchymal stem cell    microvesicles for treatment of Escherichia coli endotoxin-induced    acute lung injury in mice. Stem Cells. January 2014; 32(1):116-125.-   50. Zhang Y, Chopp M, Meng Y, et al. Effect of exosomes derived from    multipluripotent mesenchymal stromal cells on functional recovery    and neurovascular plasticity in rats after traumatic brain injury. J    Neurosurg. April 2015:122(4):856-867.-   51. Lombardo E, van der Poll T, DelaRosa O, Dalemans W. Mesenchymal    stem cells as a therapeutic tool to treat sepsis. World J Stem    Cells. Mar. 26 2015; 7(2):368-379.-   52. Yao Y, Zhang F, Wang L, et al. Lipopolysaccharide    preconditioning enhances the efficacy of mesenchymal stem cells    transplantation in a rat model of acute myocardial infarction. J    Biomed Sci. Aug. 20 2009; 16:74.-   53. Roszer T. Understanding the Mysterious M2 Macrophage through    Activation Markers and Effector Mechanisms. Mediators Inflamm. 2015;    2015:816460.-   54. Meisel R, Zibert A, Laryea M, Gobel U, Daubener W, Dilloo D.    Human bone marrow stromal cells inhibit allogeneic T-cell responses    by indoleamine 2,3-dioxygenase-mediated tryptophan degradation.    Blood. Jun. 15 2004; 103(12):4619-4621.-   55. Fernando M R, Reyes J L, Iannuzzi J, Leung G, McKay D M. The    pro-inflammatory cytokine, interleukin-6, enhances the polarization    of alternatively activated macrophages. PLoS One. 2014; 9(4):e94188.-   56. Choi J S, Kim K H, Lau L F. The matricellular protein CCN1    promotes mucosal healing in murine colitis through IL-6. Mucosal    Immunol. November 2015; 8(6):1285-1296.-   57. Kondo M, Yamaoka K, Sakata K, et al. Contribution of the    Interleukin-6/STAT-3 Signaling Pathway to Chondrogenic    Differentiation of Human Mesenchymal Stem Cells. Arthritis    Rheumatol. May 2015; 67(5):1250-1260.-   58. Hallahan D E, Virudachalam S. Intercellular adhesion molecule 1    knockout abrogates radiation induced pulmonary inflammation. Proc    Natl Acad Sci USA. Jun. 10 1997; 94(12):6432-6437.-   59. Chow A, Huggins M, Ahmed J, et al. CD169(+) macrophages provide    a niche promoting erythropoiesis under homeostasis and stress. Nat    Med. April 2013; 19(4):429-436.-   60. Jacobsen R N, Perkins A C, Levesque J P. Macrophages and    regulation of erythropoiesis. Curr Opin Hematol. May 2015;    22(3):212-219.-   61. Wang Z, Xu L, Hu Y, et al. miRNA let-7b modulates macrophage    polarization and enhances tumor-associated macrophages to promote    angiogenesis and mobility in prostate cancer. Sci Rep. May 9 2016;    6:25602.

We claim:
 1. A method for generating an educated macrophage, the methodcomprising the steps of: isolating extracellular vesicles from amesenchymal stem cell previously exposed to lipopolysaccharide (LPS),and co-culturing a CD14+ cell with the extracellular vesicles in vitrountil the CD14+ cell acquires an anti-inflammatory macrophage phenotype.2. The method of claim 1, wherein the CD14+ cell and the extracellularvesicles are co-cultured for at least 2 days.
 3. The method of claim 1,wherein the mesenchymal stem cell was exposed to LPS for at least 2hours.
 4. The method of claim 1, wherein the mesenchymal stem cell isexposed to about 50 ng/ml to about 200 ng/ml LPS.
 5. A population ofanti-inflammatory macrophages produced by the method of claim 4, whereinthe anti-inflammatory macrophage phenotype is characterized as CD206high, PD-L1 high, PD-L2 high, CD16 high and CD73 high compared tocontrol macrophages.
 6. The method of claim 1, wherein the CD14+ cell isa macrophage.
 7. The method of claim 1, wherein the CD14+ cell is amonocyte and wherein the CD14+ monocyte and the extracellular vesicleare co-cultured for at least 5 days.